INTERACTIONS BETWEEN HOST and PHAGE ENCODED FACTORS SHAPE PHAGE INFECTION a Dissertation by DENISH KUMAR PIYA Submitted To

INTERACTIONS BETWEEN HOST AND PHAGE ENCODED FACTORS SHAPE

PHAGE INFECTION

A Dissertation

DENISH KUMAR PIYA

Submitted to the Office of Graduate and Professional Studies of Texas A&M University in partial fulfillment of the requirements for the degree of

DOCTOR OF PHILOSOPHY

Chair of Committee, Ryland F. Young Co-Chair of Committee, Jason J. Gill Committee Members, Jennifer Herman Lanying Zeng Head of Department, Gregory D. Reinhart

December 2018

Major Subject: Biochemistry

ABSTRACT

Evolution of phages and their bacterial hosts are directed by interaction between phage and host- encoded factors. These interactions have resulted in the development of several defense and counter-defense strategies such as DNA restriction and antirestriction systems. Type I restriction- modification (R-M) systems present a barrier to foreign DNA, including phage, entering the bacterial cell, by cleaving inappropriately modified DNA in a sequence-specific manner. Phages have evolved diverse mechanisms to overcome restriction systems. The temperate coliphage P1 encodes virion-associated proteins that protect its DNA from host type I R-M systems. By using genetic and biochemical analysis, it has been established that the P1 Dar (Dar for defense against restriction) system is comprised of at least six virion-associated proteins: DarB, Ulx, Hdf, DarA,

DdrA and DdrB. DarB protects P1 DNA from EcoB and EcoK restriction in cis by an unknown mechanism and is incorporated into the virions only in the presence of Hdf, DarA and DdrA. Hdf and DarA have also been found to affect capsid morphogenesis, as their absence results in phage progeny with predominantly aberrant small heads. Examination of purified P1 proheads shows that Dar system proteins are incorporated into the virion before DNA packaging, and an N- terminal signal is required for DarB packaging. Twenty-four additional P1 genes of unknown function were disrupted and none were found to alter the antirestriction phenotype. A purification protocol for the ~250 kDa antirestriction protein DarB has been optimized, which will facilitate biochemical approaches for determining its mechanism of action.

While the phage-host interactions of classical phages such as P1 are better understood, relatively little is known about how phages outside of the common paradigm organisms interact with their ii

hosts. A high throughput genetic screen of the T1-like coliphage LL5 and the rV5-like coliphage

LL12 was conducted against the E. coli Keio collection. This screen revealed host receptors required for both phages to initiate infection and two chaperones, PpiB and SecB, required for efficient propagation of phage LL5.

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DEDICATION

I would like to dedicate this work to my late grandfather. He was my first teacher and taught me to read and write, and always encouraged me to pursue higher education.

ACKNOWLEDGEMENTS

I would like to thank my advisers, Dr. Jason J. Gill and Dr. Ry Young for believing in me and providing me with opportunity to work under their tutelage. Their support and words of wisdom have been very crucial in my journey through graduate school. The work presented in this dissertation would not have possible without their mentorship. Both of you have greatly inspired me to pursue science and excellence. I would also like to thank my committee members, Dr.

Jennifer Herman and Dr. Lanying Zeng for their guidance, helpful discussion and comments throughout graduate school.

I would like to thank past and present members of Gill Lab (Jackie Grimm, Season Xie, Abby

Korn, Dr. Justin Leavitt, Justin Boeckman, Miguel F. Gonzales) for creating an awesome Lab environment and always being there for helpful discussions and assistance. I would also like to thank my undergraduate research mentees Brian Koehler, Judith Salazar, Leonardo Vara and

Ashley Stonecipher for their help in conducting experiments and CPT staff Lauren Lessor for providing phages LL5 and LL12 used in this study. I also thank Dr. Matthew Taylor from

Department of Animal Science at Texas A&M University for providing STEC strains used in this study. I also thank Dr. Junjie Zhang and Dr. Timothy Devarenne from Department of

Biochemistry & Biophysics and Dr. Carlos F. Gonzalez from Department of Plant Pathology and

Microbiology at Texas A&M University for allowing me to use their lab equipment. I also thank

Dr. William Russell from Department of Chemistry at Texas A&M University (Currently at The

University of Texas Medical Branch at Galveston) and Dr. Lawrence Dangott from Department of Biochemistry & Biophysics at Texas A&M University for assistance with mass spectrometry. v

I would like to thank all CPT members and Supergroup members for their feedback and comments on data and presentations. Special thanks to Adriana C. Hernandez, Dr. Hem Thapa,

Dr. Karthik Chamakura and Dr. Manoj Rajaure for teaching me several experimental techniques.

I would also like to thank all administrative staffs of TAMU BioBio department for their assistance. Special mention to Mrs. Daisy Wilbert for always taking care of everything.

I also thank my teachers, instructors and professors from Notre Dame School (Bandipur, Nepal),

Small Heaven School (Narayanghat, Nepal), Budhanilkantha School (Kathmandu, Nepal) and

Caldwell University (Caldwell, NJ, U.S.) for providing me with a solid foundation required for this journey. I would also like to thank all of my friends from my schools in Nepal and especially from Caldwell University and Texas A&M University for making life more fun and for being there at tough times.

Finally, thanks for my parents for their encouragement and support throughout my educational journey. This journey has been only possible because their constant motivation since my first day at preliminary school. Thanks for my sisters as well for their support and love. Also, thanks to my wife for her patience and love and for sharing this journey with me.

CONTRIBUTORS AND FUNDING SOURCES

This work was supervised by a dissertation committee consisting of Dr. Ry Young (advisor), Dr.

Jennifer Herman and Dr. Lanying Zeng of the Department of Biochemistry & Biophysics and

Dr. Jason J. Gill (co-advisor) of the Department of Animal Science. All work for the dissertation was completed independently by the student. Graduate study was supported by start-up funding from Texas AgriLife Research and Texas A&M University to Dr. Jason J. Gill and from

Deerland Enzymes.

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NOMENCLATURE

E. coli Escherichia coli

R-M Restriction-Modification

viii

TABLE OF CONTENTS

Page

ABSTRACT ...... ii

DEDICATION ...... iv

ACKNOWLEDGEMENTS ...... v

CONTRIBUTORS AND FUNDING SOURCES ...... vii

NOMENCLATURE ...... viii

TABLE OF CONTENTS ...... ix

LIST OF FIGURES ...... xi

LIST OF TABLES ...... xiii

CHAPTER I INTRODUCTION ...... 1

Bacteriophages ...... 1 Phage P1 ...... 6 Host defense against phages ...... 13 Assembly of capsid-associated proteins ...... 31 Questions to be addressed ...... 33

CHAPTER II THE MULTICOMPONENT ANTIRESTRICTION SYSTEM OF PHAGE P1 IS LINKED TO CAPSID MORPHOGENESIS ...... 37

Introduction ...... 37 Materials and methods ...... 42 Results and discussion ...... 52 Conclusions ...... 77

CHAPTER III CHARACTERIZATION OF P1 GENES OF UNKNOWN FUNCTION AND PACKAGING OF DARB INTO P1 PROCAPSIDS ...... 83

Introduction ...... 83 Materials and methods ...... 86 Results and discussion ...... 105 Conclusions ...... 126 ix

CHAPTER IV GENOME-WIDE SCREENS REVEAL ESCHERICHIA COLI GENES REQUIRED FOR GROWTH OF T1-LIKE PHAGE LL5 AND RV5-LIKE PHAGE LL12 ...... 135

Introduction ...... 135 Materials and methods ...... 139 Results and discussion ...... 144 Conclusions ...... 184

CHAPTER V CONCLUSIONS AND FUTURE DIRECTIONS ...... 187

REFERENCES ...... 192

LIST OF FIGURES

FIGURE Page

1.1 Morphotypes of Caudovirales phages ...... 2

1.2 General assembly pathway of Caudovirales phages ...... 4

1.3 Lytic vs lysogenic lifecycle ...... 6

1.4 Lysis-lysogeny decision in P1 ...... 11

1.5 Host defense against different stages of phage infection cycle ...... 14

1.6 Fate of foreign DNA depends on the methylation status of the recognition sequence ...20

1.7 Unmethylated phage DNA is cleaved by host type I R-M system ...... 24

2.1 The antirestriction system of phage P1 protects DNA from type I restriction and modification (R-M) mediated cleavage ...... 38

2.2 darB and ulx are required for protection of P1 against EcoB and EcoK restriction ...... 55

2.3 hdf, darA and ddrA are required for protection of P1 against EcoA, EcoB and EcoK restriction ...... 57

2.4 ddrB negatively affects the darB phenotype ...... 60

2.5 Comparative SDS-PAGE of parental P1 and isogenic mutants suggests cascading dependencies for protein incorporation into the P1 virion ...... 62

2.6 Identification of Ulx band in SDS-PAGE of P1 virions ...... 64

2.7 Purification of virions using Cesium chloride isopycnic gradient centrifugation ...... 69

2.8 Head size variation in parental P1 and mutants ...... 71

2.9 Comparative SDS-PAGE of P1N and P1S virions ...... 76

2.10 Models of P1 head-size determination and antirestriction component incorporation ...... 78

3.1 Genome map of P1 ...... 85

3.2 P1 genes of unknown function are not required for antirestriction ...... 108

FIGURE Page

3.3 P1 antirestriction components are incorporated into P1 procapsids before DNA ...... 111

3.4 DarB is synthesized in P1ΔdarA mutants ...... 114

3.5 P1 procapsids show electron densities suggesting presence of protein contents ...... 115

3.6 Plating efficiency of P1ΔdarB complemented with truncated DarB ...... 119

3.7 Truncated DarB can be expressed ...... 121

3.8 N-terminal residues of DarB provide signal for capsid targeting ...... 122

3.9 IMAC purification of P1 DarB ...... 127

3.10 DarB with 2x His-tag fused to N-terminus is functional ...... 128

4.1 Transmission electron micrographs of phages LL5 and LL12 ...... 145

4.2 Genome maps of phages LL5 and LL12 ...... 147

4.3 Genes and biosynthetic pathway of the E. coli lipopolysaccharide (LPS) required for replication of phages LL5 and LL12 ...... 179

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LIST OF TABLES

TABLE Page

1.1 Characteristics of different R-M systems ...... 21

1.2 Antirestriction mechanism of phages ...... 25

1.3 Recognition sequence of type I R-M systems ...... 29

2.1 Bacterial strains, phages and plasmids ...... 43

2.2 Primers ...... 45

2.3 Copy number of virion-associated proteins of P1 antirestriction system ...... 68

2.4 Mutations in phage P1 antirestriction genes affect head morphogenesis ...... 73

3.1 Bacterial strains, phages and plasmids ...... 87

3.2 Primers and synthetic DNA fragments ...... 91

4.1 Proteins encoded by genome of phage LL5 ...... 149

4.2 Proteins encoded by genome of phage LL12 ...... 153

4.3 Analysis of LL12 tail fibers ...... 162

4.4 Host range of phages LL5 and LL12 ...... 166

4.5 Summary of hits in Keio screening ...... 168

4.6 Results of initial (untargeted) screening and targeted re-screening of phage LL5 against the Keio E. coli knockout collection ...... 169

4.7 Results of initial (untargeted) screening and targeted re-screening of phage LL12 against the Keio E. coli knockout collection ...... 172

4.8 E. coli genes required for phages LL5 and LL12 propagation ...... 177

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CHAPTER I

INTRODUCTION

Bacteriophages

Bacteriophages (phages) are natural predators of bacteria. It has been estimated that

~96% of phages existing in nature belong to order Caudovirales, characterized by the presence of double stranded DNA as genomic material in the capsids and tails as infection apparatus (Brussow & Hendrix, 2002; Hatfull & Hendrix, 2011). Tailed phages are the most abundant life forms as they have been estimated to be present in numbers

>1030 and outnumber their bacterial hosts by five- to ten-fold in environment (Brussow

& Hendrix, 2002; Hendrix, 2002). Phages are categorized into different taxonomic groups based upon a system proposed by the International Committee on Taxonomy of

Viruses (ICTV) (Nelson, 2004). Phages of order Caudovirales are further classified, based on tail morphology into different families: Myoviridae (long, straight, contractile tail), Siphoviridae (long, flexible, non-contractile tail) and Podoviridae (short, stubby, noncontractile tail) (Fig. 1.1) (Maniloff & Ackermann, 1998). This system of classification, based on shared characteristics, however, has various limitations. This system does not consider the genomic and proteomic information, resulting in resulting in grouping of unrelated phages based on their morphology only (Botstein &

Herskowitz, 1974; Nelson, 2004). Since phage replication involves genetic recombination, phage genomes are mosaic, which is not considered in this classification system. It is also not feasible to isolate phages for determining its morphological

characteristics when the phage genomes are obtained from community sequencing project (Nelson, 2004).

Advances in structural biology has made analysis of phage structure in atomic resolution possible. Based on the conserved folds of structural proteins, Caudovirales phages are suggested to have a common evolutionary origin. The assembly of these virions occur

A B C

Fig. 1.1. Morphotypes of Caudovirales phages. Phages belonging to order Caudovirales are divided into three families, based upon characteristics of their tails. Panel A: Phages of family Myoviridae have long contractile tail. Panel B: Phages of family Siphoviridae have long, noncontractile but flexible tails. Panel C: Phages of family Podoviridae have short tails.

from conserved components in a similar pathway (Fig. 1.2) (Fokine & Rossmann, 2014).

In general, the capsids are icosahedral and are composed of eleven pentameric, and twenty hexameric capsomers (Mateu, 2013). One pentameric vertex is occupied by a dodecameric ring of portal protein and is called the portal vertex (Aksyuk & Rossmann,

2011). The assembly of capsids start at the portal vertex in presence of scaffolding proteins and major capsid proteins, forming proheads (Fig. 1.2). The scaffolding proteins facilitate the assembly of capsid proteins into a definite geometry (Dokland, 1999). DNA is packaged into procapsids through the ring of portal proteins by DNA packaging machine composed of small and large terminases (Casjens, 2011). Once the DNA is packaged, the terminase complex disassociates and the head completion proteins are attached to the portal vertex. The head completion proteins serve as the attachment site for tails of Myoviridae and Siphoviridae phages, which are assembled in an independent pathway. The assembly of tails in Myoviridae and Siphoviridae start at the base plate and the hexamers of tail tube and tail sheath are sequentially added to a certain length as determined by tape measure protein. However, in Podoviridae phages, tails are assembled on the portal vertex (Aksyuk & Rossmann, 2011; Fokine & Rossmann, 2014).

Fig. 1.2. General assembly pathway of Caudovirales phages. Schematics represent assembly pathway of Myoviridae and Siphoviridae phages. Major capsid proteins, scaffolding core and portal proteins assemble into procapsids. DNA is packaged into procapsids by DNA packaging complex, followed by maturation of capsids into icosahedral geometry. Tails are assembled in an independent pathway and are attached to DNA-filled capsids to form complete virions. In Podoviridae phages, tails are assembled directly into the portal vertex.

Phage infection cycle starts with adsorption of phages to the specific receptors on cell surface of a host, mediated by the phage receptor-binding protein, also called tail fiber proteins (Dowah & Clokie, 2018). This adsorption event is followed by ejection of phage genetic material into host cytoplasm. Depending upon if the infecting phage is virulent or temperate, a specific life cycle is pursued (Fig. 1.3) (Gill & Hyman, 2010).

Virulent phages always pursue lytic life cycle, which begins with sequential expression of phage proteins to take over host cell, which usually follows by DNA replication and synthesis of structural components. Complete virions are assembled and host cells are lysed to liberate progeny phages to surroundings (Echols, 1972). Contrary to the lifecycle of virulent phages, temperate phages can pursue lytic or lysogenic life cycle.

The lytic cycle is pursued in a similar fashion as described for virulent phages, whereas in lysogenic cycle, phage genome is integrated into host chromosome, in most cases and phage-host genome replicate simultaneously. During lysogenic state, genes responsible for synthesis of structural proteins and host lysis are repressed. Under certain conditions, lysogenic state can be induced to lytic cycle and phage progeny are released into surroundings upon host lysis (Echols, 1972).

A B

Fig. 1.3. Lytic vs lysogenic life cycle. Phage infection cycle starts with adsorption of a phage to its host receptors, followed by ejection of its DNA into host cytoplasm.

Depending upon whether infecting phages are virulent or temperate, phages can pursue either lytic or lysogenic life cycle. Panel A: In lytic life cycle, phage DNA replicates and simultaneously various components necessary for virion morphogenesis are synthesized. Phage DNA is packaged into procapsids by DNA packaging complex. The phage capsids and tails are attached forming mature virions. At a genetically determined time, the host cell lyses releasing progeny virions into the surroundings.

Panel B: In lysogenic life cycle, phage DNA is mostly integrated into host chromosome. Phage DNA and host DNA are replicated simultaneously. Virulent phages can pursue only lytic life cycle, whereas temperate phages can pursue either lytic or lysogenic life cycle. Temperate phages can also switch to lytic life cycle from lysogenic state.

Phage P1

Phage P1 was discovered as a resident prophage in E. coli strain “Li” , along with two other phages, P2 and P3, by G. Bertani in 1951 (Bertani, 1951). These three phages, P1,

P2, and P3, could be differentiated by plaque morphology as well as serum inactivation assay. The plaques formed by P1 were small, with size of less than 1 mm in diameter

(Bertani, 1951). Along with other E. coli phages such as , T4 and T7, P1 has played a crucial role in the development of molecular biology and the study of phage infection cycle. T4 and T7 have a virulent lifecycle whereas both  and P1 have temperate lifecycles (Lobocka et al., 2004). Unlike phage  that is integrated into host chromosome during lysogeny, P1 is maintained as an extra-chromosomal plasmid (Ikeda

& Tomizawa, 1968). P1 has been used as a workhorse of gene transduction as it can non-specifically package host chromosome (Lennox, 1955; Lobocka et al., 2004).

Virion morphology

P1 virions have been analyzed extensively by negative-stained electron microscopy. P1 is a member of Myoviridae family and has an icosahedral head, and a long contractile tail that terminates in six tail fibers (Walker & Anderson, 1970; Lobocka et al., 2004). It has been reported that the lysates of P1 consist of virions of three different head sizes:

P1B with ~86 nm, P1S with ~65 nm, and P1M with ~45 nm side-to-side diameter. The ratio of sizes of P1B:P1M:P1S is 4:3:2, which corresponds to T values of 16, 9 and 4 for

P1B, P1M and P1S respectively (Walker & Anderson, 1970). Only the virions of head- size class P1B can package the full-length genome, hence are infectious. On the contrary, virions of both P1S and P1M cannot package full-length genome and thus, do not have a complete set of genetic elements required to establish a successful infection

(Walker & Anderson, 1970).

The host specificity of P1 is determined by a 4.2 kb invertible C segment which involves a similar mechanism as in phage Mu (Iida, 1984). The C segment consists of tail fiber genes Sv-U or Sv’-U’, encoding the variable region of tail fibers, which is fused to the constant region encoded by Sc (Guidolin et al., 1989a). Cin, a site-specific recombinase, is encoded by a gene adjacent to the C segment and mediates recombination between the

0.6 kb inverted repeats flanking the C segment, which form the cix ( for C inversion cross-over) sites (Iida et al., 1982). P1 virions induced from lysogens have both C(+) and C(-) orientations, whereas P1 virions produced from lytic infection have only C(+) orientation (Iida et al., 1982). P1 with C(+) orientation can infect both E. coli K12 and E. coli C, whereas P1 with C(-) orientation only infects an E. coli mutant (Iida et al., 1982).

DNA packaging

The infective virions of P1 contain of ~94 kb cyclically permuted, linear, double- stranded DNA with a terminal redundancy of 10-15 kb (Sternberg, 1990; Lobocka et al.,

2004). P1 genome consists of 117 predicted genes, organized into 45 operons, of which

112 are translated into proteins and five code for untranslated RNA’s (Lobocka et al.,

2004). The genomic DNA is produced as linear concatemers and packaged into P1 procapsids by a headful mechanism by the P1-encoded pacase proteins, PacA and PacB

(Skorupski et al., 1994a). P1 DNA contains a consensus pac sequence, which is recognized and cleaved by P1 pacase proteins, followed by packaging of the cleaved

DNA into P1 procapsids. After the first headful packaging, the pacase proteins cleave

DNA in a non-sequence-dependent manner and initiates packaging of the remaining linear concatemeric DNA substrate into a new procapsid. After DNA packaging, P1 tails and DNA-filled capsids are attached to form complete P1 virions (Sternberg, 1990;

Skorupski et al., 1994b).

Cyclization of P1 DNA following infection

After recognizing a specific bacterial host, P1 ejects its DNA into host cytoplasm. Once in the cytoplasm, unlike other phage DNA, P1 DNA exists as a self-replicating, extrachromosomal, circular form. P1 encoded recombinase protein Cre (for causes recombination) and loxP [for locus of crossing over (x), P1] sites, at which recombination occurs, are necessary for circularization of P1 DNA following infection

(Sternberg & Hamilton, 1981; Hochman et al., 1983). The conversion of the linear DNA in the P1 capsid to the covalently closed circular form in host cytoplasm, after infection, occurs by an intramolecular recombination event that leads to the removal of the redundant portion of the DNA (Segev et al., 1980). This recombination event results in the genetic map of P1 being linear, despite P1 DNA being cyclically permuted, in contrast to the circular genetic maps of other phages such as P22 or T4 (Sternberg &

Hamilton, 1981). Because of the linearity of P1 genome, P1 genes on the ends of the genome are not linked together (Sternberg & Hamilton, 1981). The Cre-loxP system also mediates the segregation of P1 prophage into daughter host cells during lysogenic growth by resolving a P1 dimer composed of two P1 monomers (Sternberg et al., 1986).

After it was discovered that this Cre-loxP system can function in yeast Saccharomyces

cerevisiae, it has been used extensively in other eukaryotes for genome editing (Sauer,

1987; Lambert et al., 2007). P1 DNA is also circularized by the host RecBCD nuclease complex. There is a total of 50 recombinational host spot Chi sites in positive and negative strands of P1 DNA. RecBCD complex functions as a 3’-to-5’ nuclease, but when this complex encounters Chi sites in P1 genome, the enzymatic activity of the complex is switched to recombination-promoting form and thus helps in homologous recombination mediated circularization of P1 DNA (Zabrovitz et al., 1977; Lobocka et al., 2004).

Lysis-lysogeny decision

After circularization of DNA, P1, being a temperate phage, can pursue either lytic or lysogenic infection cycle. This lytic-lysogenic decision is controlled by genes in P1 immunity circuitry, which is distributed in three regions of P1 genome: ImmC, ImmI and

ImmT (Fig. 1.4) (Yarmolinsky, 2004). Among the temperate phages studied, the immunity region of P1 and P7 is the most complex one (Heinrich et al., 1995). The

ImmC region consists of the c1 (master repressor) and coi (C1 inhibitor). The ImmI region include genes that code for translational repressor RNA, C4, and antirepressor proteins, Ant1and Ant2. The ImmT region has a gene, lxc that codes for a corepressor

(Fig. 1.4) (Yarmolinsky, 2004). C1 and Coi are antagonistic to each other (Yarmolinsky,

2004). C1 repressor binds to ~17 operators throughout P1 genome and negatively

Fig. 1.4. Lysis-lysogeny decision in P1. Panel A: The decision to pursue lytic or lysogenic lifecycle in P1 is determined by the immunity circuitry, located in three regions of temperate phage P1 genome (represented by circle): ImmC, ImmT and ImmI. ImmC encodes C1 (master repressor) and Coi, ImmT encodes Lxc and ImmI encodes C4 (RNA), Ant1 and Ant2. Panel B: The lytic or lysogenic lifecycle is decided by the interplay between different factors. If C1 prevails, lysogeny is established, whereas if Coi prevails, lytic replication is established (Heinrich et al., 1995), (Yarmolinsky, 2004), (Lobocka et al., 2004).

regulates the expression of genes required for lytic pathway, including Coi, whereas Coi inactivates C1 by direct interaction. The RNA, C4 blocks the ribosome binding site in the transcript of the antirepressor proteins, thereby reducing the cellular concentration of the antirepressor proteins. These antirepressor proteins inactivate the C1 repressor

(Heinrich et al., 1995; Yarmolinsky, 2004). Lxc forms a complex with C1, and enhances the binding affinity of C1 to its operators, consequently resulting in the lowered expression of C1 itself (Yarmolinsky, 2004). Moreover, the ability of Coi to inhibit C1 repressor activity is inhibited in the presence of Lxc. The decision to pursue lytic or lysogenic lifecycle, upon infection of host by P1, is determined by the competition between C1 and Coi proteins (Heinrich et al., 1995). If C1 synthesis is maintained, lysogeny growth is established, whereas if Coi synthesis is maintained, lytic growth is established (Lobocka et al., 2004). In our study, we have used P1 with temperature sensitive C1 repressor. At non-permissive temperature, C1 is degraded and lytic infection cycle is induced (Lobocka et al., 2004).

Host lysis

Phages of Gram-negative bacterial hosts are released into the surroundings after the disruption of the cellular membrane and the peptidoglycan layer. Usually, host lysis is accomplished by actions of three proteins. The holins create “holes” in the inner membrane releasing endolysins to the periplasmic space where the peptidoglycan layer is degraded by enzymatic activity of the endolysins (Young, 2013). In the final step, spanin complexes fuse the inner and the outer membrane resulting in the host lysis

(Rajaure et al., 2015). The host lysis steps mediated by P1 is unique in that the export of endolysins to the periplasmic space is not dependent on holins (Xu et al., 2004). P1 endolysin, Lyz, has a N-terminal SAR (for signal-anchor and release) sequence and is exported and tethered in an inactive form to the inner membrane by the sec translocon.

The SAR sequence is also sufficient for the release of Lyz from the inner membrane into the periplasmic space, without any proteolytic cleavage of the sequence, and this release occurs because of the collapse of the membrane potential(Xu et al., 2004).

Host defense against phages

It has been estimated that ~1023 phage infections occur every second (Hatfull & Hendrix,

2011), which provide enormous selection pressure on bacterial host. Since bacteria need to survive constant predation by phages, bacteria have evolved a wide arsenal of defense systems to thwart the phage infection cycle (Fig. 1.5). In turn, phages are constantly evolving with antidefense systems to overcome these bacterial defense systems. This constant co-evolution of phages and their bacterial hosts have led to an evolutionary arms race, resulting in significant diversity in defense systems of bacteria and phages against each other (Van Valen, 1973; Labrie et al., 2010; Stern & Sorek, 2011).

Fig. 1.5. Host defense against different stages of phage infection cycle. Bacteria have defense mechanisms to thwart phage infection at various stages. Bacteria can prevent adsorption of phages to the receptors by masking receptors with extracellular materials or by modifying receptors themselves. Even after successful adsorption, bacteria can prevent the ejection of phage DNA into cytoplasm by obstructing the DNA entry point. In the cytoplasm, phage DNA can be cleaved by CRISPR-Cas systems or Restriction- Modification systems. Moreover, bacteria possess abortive infection or assembly interference mechanisms that eventually prevent the formation of mature virions (Murray, 2000; Labrie et al., 2010; Seed, 2015). The image is adapted from Seed, 2015.

Changes in receptors

The first step of phage infection cycle is the recognition and adsorption to the specific receptors on the surface of host cell by the receptor binding proteins of the phages. The receptors are present on the outer surface of the host cells and may be comprised of proteins, polysaccharides and lipopolysaccharides (Samson et al., 2013). Bacteria have different strategies to prevent the adsorption of phages. The common theme of these strategies is to mask the specific receptor component. In Staphylococcus aureus, increased secretion of protein A has been shown to mask specific phage receptors

(Nordstrom & Forsgren, 1974). Similarly, the K1 polysaccharide capsule of E. coli has been shown to prevent T7 infection (Scholl et al., 2005). In another instance, in

Bordetella spp., phase variation, under the control of the BvgAS two-component regulatory system, determines the expression of receptors and hence, phage infection

(Liu et al., 2002). Similar phase variation has also been reported in Vibrio cholerae O1 serogroup strains, in which the expression of O1 antigen determines if the cells are susceptible to phage infection or not (Seed et al., 2012). Likewise, in other cases, extracellular components such as alginates, hyaluronan and glycoconjugates act as a barrier to prevent phages from accessing their specific receptors (Labrie et al., 2010).

Phages have evolved with strategies to overcome this barrier provided by bacterial cells which masks receptors. By generating mutations in the host specificity determining gene mtd, Bordetella phage BPP-1 can infect host expressing different receptors (Liu et al.,

2002). Besides, Pseudomonas phage F116 and AF has been shown to degrade the

exopolysaccharide matrix surrounding the host, which might be necessary to reach the cell surface and initiate infection (Hanlon et al., 2001; Cornelissen et al., 2012).

Moreover, several instances have also been reported where phages have acquired the ability to recognize these extracellular barriers as host receptors (Gross et al., 1977;

Steinbacher et al., 1997).

Blocking DNA entry into cytoplasm

Only after recognition of the cell surface receptors, phage genetic material is ejected into host. This stage of phage infection cycle can be prevented by superinfection exclusion

(Sie) systems. The Sie systems are encoded by phage genomes; thus, these systems specifically define the phage defense systems to prevent infection by closely related phages. The Sie proteins are localized in the membranes and prevent the phage genetic material from being delivered to the cytoplasm (Labrie et al., 2010). The well-studied coliphage T4 has two Sie proteins, Imm and Sp. Imm prevents the transfer of T4 DNA into the cytoplasm by interfering with the injection site (Vallee & Cornett, 1972; Lu &

Henning, 1989). T4 tail-associated protein gp5 assists in DNA ejection process with its lysozymic activity by degrading peptidoglycan layer. The other Sie protein, Sp inhibits the activity of T4 gp5, thereby blocking DNA ejection into cytoplasm (Kao & McClain,

1980). Other Sie systems are also described in coliphage P1 (Kliem & Dreiseikelmann,

1989), HK97 (Cumby et al., 2012) and Salmonella enterica phage P22 (Susskind et al.,

1971; Susskind et al., 1974a; Susskind et al., 1974b).

The Sie system of P1 is encoded by sim gene in immC region of P1 genome (Kliem &

Dreiseikelmann, 1989). Sim is synthesized as a ~25 kDa protein, which is proteolytically processed at its N-terminus to yield a ~24 kDa protein in a SecA-dependent manner. Sim is predicted to localize either in the membrane or the periplasmic space and prevents P1 superinfection (Maillou & Dreiseikelmann, 1990).

Degradation of phage DNA

Once successfully introduced to the host cytoplasm, the phage genetic material still must overcome host defenses such as CRISPR (for Clustered regularly interspaced short palindromic repeats)- Cas (for CRISPR-associated) and R-M (for Restriction-

Modification) systems mediated cleavage of the genetic material (Labrie et al., 2010).

CRISPR-Cas

CRISPR-Cas systems are the nucleic-acid based adaptive immunity systems that provide protection against foreign DNA including phage DNA, in which DNA fragments of an infecting phage are incorporated into CRISPR arrays and subsequent infection from the same phage is prevented (Barrangou et al., 2007). CRISPR arrays consist of several partially palindromic repeats (CRISPR repeats, 23-55 nt) separated by variable (CRISPR spacers, 21-72 nt). These CRISPR arrays are often adjacent to cas genes (Barrangou et al., 2007; Barrangou & Marraffini, 2014). In general, protection provided by CRISPR-

Cas systems involve three distinct steps. In the first step, the spacer sequences are acquired from the genome of the phages that have infected the host. In the next step, the

CRISPR arrays are transcribed and processed into CRISPR RNAs, which form a complex with Cas proteins. Eventually, upon infection by the same phage, the complementary base-pairing of CRISPR RNAs and phage genome results in the degradation of the target (Stern & Sorek, 2011; Barrangou & Marraffini, 2014; Leon et al., 2018). CRISPR-Cas systems are categorized into two major classes based on the organization of effector proteins, that mediate CRISPR RNA processing, and target recognition and cleavage (Koonin et al., 2017).

Phage have diverse mechanisms to prevent CRISPR-Cas mediated DNA degradation.

Any mutations in the spacer targeted (protospacer) region or the PAM (for protospacer- adjacent motif) will render CRISPR-Cas ineffective (Deveau et al., 2008; Pawluk et al.,

2018). Moreover, recent studies have discovered several anti-CRISPR proteins encoded in phage genomes, that inhibit CRISPR-Cas systems (Bondy-Denomy et al., 2013;

Pawluk et al., 2016a; Pawluk et al., 2016b; Rauch et al., 2017).

Restriction-Modification systems

The R-M systems are the innate host defense systems which specialize in cleaving the foreign DNA based on its modification status. The effects of R-M systems were initially observed as the host specificity imparted to phages after they were propagated on different hosts (Bertani & Weigle, 1953). It was later demonstrated that the factors determining host specificity were physically linked to phage DNA (Arber & Dussoix,

1962) and degradation of phage DNA resulted in the host specificity of phages (Dussoix

& Arber, 1962). The R-M systems are broadly categorized into three types, type I, II and

III, based on subunit composition, cofactor requirements, recognition site and cleavage position (Murray, 2000; Tock & Dryden, 2005). This study focuses on the P1 components that are required to protect phage DNA from the restriction function of type

I R-M system.

The genes encoding components of type I R-M system were first identified in 1970

(Hubacek & Glover, 1970). The type I R-M systems are hetero-oligomeric complexes, encoded by three genes hsdS, hsdM and hsdR (hsd for host specificity determinant) (Fig.

1.6) (Murray, 2000). Unlike type I R-M systems, type II and III are composed of two subunits only. The main characteristics of these two systems and their differences with type I R-M systems are highlighted in (Table 1.1). HsdS (S) is the specificity subunit that enables any type I R-M complex to recognize specific base sequences. HsdM (M) is the modification subunit that adds methyl group in a sequence-dependent manner and

HsdR (R) is the restriction subunit that cleaves the DNA under certain conditions (Tock

& Dryden, 2005). The three subunits of type I R-M system can form two forms of complexes; modification complex and restriction complex. Both complexes recognize an asymmetric, bipartite sequence. The modification complex is composed of M and S subunits in the stoichiometric ratio of 2:1 (M2S1), whereas the restriction complex is composed of all R, M and S subunits in the stoichiometric ratio of 2:2:1 (R2M2S1). The modification complex can only modify the DNA in a sequence-dependent manner, whereas the restriction complex can either modify or cleave DNA depending upon the

Fig. 1.6. Fate of foreign DNA depends on the methylation status of the recognition sequence. Panel A: The type I R-M system is a complex composed of three subunits:

HsdS (S), HsdM (M) and HsdR (R). The S subunit provides sequence specificity to the complex, the M subunit has the active site to methylate specific bases within the recognition site, whereas the R subunit has the active site to cleave the DNA. The three subunits can form two stoichiometric complexes: M2S1 and R2M2S1. M2S1 is the modification subunit that can only methylate DNA, whereas R2M2S1 can either methylate or cleave the DNA depending upon its methylation status. Panels B and C: The foreign DNA can be hemi-methylated or unmethylated (methylation status denoted by ★). (B) If the DNA is hemi-methylated , then the R2M2S1 complex, methylates the other strand. (C) If the DNA is unmethylated, the R2M2S1 complex translocates the DNA and cleaves the DNA a few kb away from the recognition site .

Table 1.1. Characteristics of different R-M systems. The characteristics of different R-M systems are highlighted (Kruger & Bickle, 1983; Tock & Dryden, 2005). The methylation activity of the R-M systems described in the table occurs with the recognition sequence, whereas the site for DNA restriction varies according to the type of R-M systems. Adenine is methylated at N6 position, whereas cytosine is methylated at either N4 or C5 position (Tock & Dryden, 2005).

DNA R-M Subunit DNA cleavage DNA Subunits Co-factors recognition Systems arrangement site translocation sequence

Modification AdoMet NA No HsdS (S), - M S 2 1 Asymmetric, type I HsdM (M), ~1-5 kb away AdoMet bipartite Restriction - from Mg2+, Yes HsdR (R) R M S recognition 2 2 1 ATP site Restriction Modification AdoMet NA No (R), - M mostly type II palindrome mostly at the Modification Restriction - Mg2+ recognition No (M) R 2 site Modification AdoMet NA No Mod (M), -M2 type III asymmetric ~25-27 bp Res (R) Restriction- Mg2+, away from Yes R M ATP 2 2 recognition site

methylation status of the recognition sequence. If the DNA is hemi-methylated, the restriction complex methylates a specific base in the complementary strand, and only if the DNA is unmodified or inappropriately modified in both strands, the restriction complex cleaves the DNA (Fig. 1.6) (Murray, 2000). The type I R-M complex requires

AdoMet, Mg2+ and ATP as cofactors. HsdM provides binding interface for AdoMet, whereas HsdR provides binding interface for Mg2+ and ATP. AdoMet acts as the methyl donor and is the only cofactor required for the modification activity. However, all of

AdoMet, Mg2+, and ATP are required for restriction activity (Table 1.1) (Meselson &

Yuan, 1968; Tock & Dryden, 2005).

Restriction of foreign DNA by type I R-M systems leading to specific activity involve a series of steps. In general, the enzyme complex gets activated by binding to AdoMet and the activated complex binds to DNA. Upon binding to ATP, the complex can recognize methylation status in recognition site. If DNA is methylated, the complex is released from DNA. If DNA is hemi-methylated, a specific base in recognition site in the complementary strand in methylated. If DNA is unmethylated, DNA is translocated and cleaved several kb away from the recognition site, accompanied by ATP hydrolysis (Fig.

1.6) (Yuan et al., 1975; Burckhardt et al., 1981).

Bacterial host may contain any specific type I R-M system. Depending upon the restriction background of the host, the DNA of progeny phage will have sequence specific modification (Fig. 1.7). On the contrary, if the bacterial host does not have any functional type I R-M system, the DNA of progeny phages will not have any sequence specific modification. If the sequence specificity of the type I R-M system of the new host is same, then the modified phage DNA, upon ejection into host cytoplasm, is not cleaved, and the phage can establish infection efficiently. In another case, if the phage

DNA is unmethylated at the recognition sequence of the type I R-M system of the new host, phage DNA is cleaved and phages have lower plating efficiency (Fig. 1.7).

Phages have evolved with diverse mechanisms to overcome the type I R-M system mediated DNA cleavage. The mechanisms of how paradigm phages such as T3, T4, T7,

Mu, , and P1 protect their genome from host type I R-M system mediated DNA cleavage have been well studied (Table 1.2 ) (Labrie et al., 2010).

Fig. 1.7. Unmethylated phage DNA is cleaved by host type I R-M system. Depending upon the restriction background of host, the phage propagated in the host could have modification in a particular recognition sequence (modification denoted by ★). Upon infection of a new host, phage DNA is encountered by the type I R-M system of the new host. If the recognition sequence of this type I R-M system in the phage DNA is modified, the DNA is not cleaved and the phage can plate efficiently. In other case, if phage DNA is unmodified at the recognition sequence, then it is cleaved and the phage cannot plate efficiently.

Table 1.2. Antirestriction mechanism of phages. Phages have diverse mechanisms of overcoming the type I R-M system of their bacterial host. Some mechanisms of well-studied phages have been mentioned in the table. See text for details.

Phages Proteins Mechanism of protection References

Dimer of Ocr mimics the phosphate backbone of (Studier & Movva, 1976; T3 and B-form DNA and binds to the type I R-M complex, Ocr Kruger & Bickle, 1983; T7 inactivating its function. T3 ocr also hydrolyzes Atanasiu et al., 2002) SAM, a co-factor for type I R-M system.

Hypermodification of DNA: The cytosine of (Volkin, 1954; Kornberg T-even N/A genome is replaced by hydroxymethyl cytosine, et al., 1961; Kruger & which is again glucosylated. Bickle, 1983)

Mu Mom Acetimidation of adenine in genome. (Kruger & Bickle, 1983)

Enhances methylation function of type I R-M (Zabeau et al., 1980;  Ral complex. Kruger & Bickle, 1983)

Proposed to methylate DNA upon ejection into host (Iida et al., 1987; Piya et P1 DarB cytoplasm. al., 2017)

Phages T3 and T7 protect DNA from type I R-M system by expressing protein gp0.3.

The protein gp0.3, also called Ocr (Ocr for overcome classical restriction), is one of the early proteins expressed following T3 and T7 infection (Studier, 1973). The dimer form of Ocr mimics the size, shape and charge of bent B-form DNA and inhibits the type I R-

M complex by directly binding to the DNA binding region of the complex (Walkinshaw et al., 2002). The binding affinity of the type I R-M complex to Ocr is stronger than that for DNA (Samson et al., 2013). Because of this direct interaction between Ocr and type I

R-M complex, the type I R-M complex cannot bind to DNA, and hence it is unable to degrade foreign DNA (Tock & Dryden, 2005). T3 Ocr protein also inhibits the type I R-

M system by hydrolyzing AdoMet, which is a co-factor required for the biological activity of type I R-M system (Studier & Movva, 1976; Spoerel et al., 1979). In the absence of AdoMet, the type I R-M system cannot be activated to degrade foreign DNA

(Tock & Dryden, 2005).

The ability of the T-even phages, T2, T4 and T6, to evade restriction provides an example of how phage-host interactions has shaped evolution. These phages incorporate an unusual base hydroxymethylcytosine (HMC) in its genomic DNA (Volkin, 1954;

Lehman & Pratt, 1960; Kornberg et al., 1961; Kruger & Bickle, 1983). Because of the presence of this unusual base, DNA of these phages are not susceptible to cleavage from

restriction complexes that have cytosine in the recognition sequence. To overcome this feature of T-even phages, host cells have acquired the modification-dependent systems

(MDSs) such as McrA, McrBC and Mrr that can cleave hydroxymethylated DNA

(Raleigh & Wilson, 1986). Interestingly, DNA of T-even phages are also glucosylated at

HMC, hence these DNAs are also protected against these MDSs (Volkin, 1954; Lehman

& Pratt, 1960; Kornberg et al., 1961; Kruger & Bickle, 1983). However, a prophage encoded two-component system consisting of proteins, GmrS and GmrD (Gmr for glucose-modified restriction), which can degrade glucosylated HMC, have been found in

E. coli strain CT596. To protect DNA from GmrS-D restriction systems, T-even phages inject hundreds of virion-associated internal protein I (IPI), which inactivate the nuclease activity of the GmrS-D restriction systems (Abremski & Black, 1979; Rifat et al., 2008;

Labrie et al., 2010).

A peculiar epigenetic modification is present in phage Mu DNA, which make its DNA insensitive to restriction from type I R-M complex because of the function of phage gene mom (mom for modification of Mu). Owing the gene function of mom, ~15% of adenine residues in Mu DNA is modified by acetimidation (Kruger & Bickle, 1983).

The genome of phage  contains the gene ral (for restriction alleviation) that contributes to protecting DNA from type I R-M systems (Zabeau et al., 1980). The antirestriction protein Ral enhances the modification activity of type I R-M complex, thereby lowering the number of the unmodified recognition sites in  genome (Zabeau et al., 1980;

Loenen & Murray, 1986; Samson et al., 2013). Because of this, cleavage of  DNA by the type I R-M complex is alleviated.

Following infection of its E. coli host, phage P1 injects Dar (for defense against restriction) proteins to protect its DNA from host type I R-M system (Iida et al., 1987).

Iida et al. discovered that two P1 loci provided restriction protection against a subset type I R-M system and hence, were named accordingly. DarA protected P1 DNA from

EcoA-mediated restrictioon, and DarB protected P1 DNA from EcoB- and EcoK- mediated DNA cleavage. Dar proteins are virion-associated and act only in cis.

However, they can protect any DNA that is packaged into P1 capsids. Packaging of

DarB into virions was known to be dependent on the presence of DarA because darA- phages were phenotypically darB- (Iida et al., 1987). DarA is synthesized as a high- molecular-weight precursor and is proteolytically processed at the N-terminus. Only the processed form of DarA is packaged into P1 virions (Streiff et al., 1987). The genes encoding these Dar proteins were found in two distinct regions of P1 genome (Lobocka et al., 2004). The genes near darA and including darA itself, were also shown to affect virion morphogenesis, but the phenotype was not distinctly assigned to any gene (Iida et al., 1998). All three of these type I R-M systems have specific recognition site and different number of those recognition sites in P1 genome (Table 1.3).

Table 1.3. Recognition sequence of type I R-M systems. Three type I R-M systems used in our study has their own sequence specificity. The recognition sequence for any type I R-M system is bipartite. There are two groups of specific sequences: One in the 5' end and the other in the 3' end. These two groups are separated by non-specific bases. The red face A (5' group) is methylated by the type I R-M complex in the positive strand, whereas the complementary base A in the to the bold face T (3' group) is methylated in the negative strand (Loenen et al., 2014). There are different number of recognition sites in the P1 genome for these type I R- M systems. Type I R-M Recognition sequence Number of sites System (5'-3') in P1 genome EcoA GAG(N7)GTCA 2 EcoB TGA(N8)TGCT 20 EcoK AAC(N6)GTGC 11

Abortive infection

Even when phages survive the R-M and CRISPR-Cas systems, other bacterial defense systems such as abortive infection and assembly interference can still prevent phage infection cycle (Chopin et al., 2005; Seed, 2015). Compared to other bacterial defense systems, abortive infection results in the death of host cell, thus this defense system is also referred to as altruistic death of host cells to prevent phage multiplication so that the surrounding bacterial population can be protected from phage predation (Fineran et al.,

2009). The phage  encoded Rex system is the best characterized abortive infection system. Upon activation, the Rex system depolarizes the membrane resulting in lowered cellular ATP concentration, which ultimately stops all cellular processes and thus prevents phage multiplication (Parma et al., 1992). Several other abortive systems have also been reported to directly target different stages of the phage infection cycle such as

DNA replication (Emond et al., 1997), phage transcripts (Parreira et al., 1996), and inducing early lysis (Durmaz & Klaenhammer, 2007).

Assembly interference

The assembly of certain phage particles can also be prevented by the action of phage- inducible chromosomal islands (PICIs) such as Staphylococcus aureus pathogenicity islands (SaPIs) (Seed, 2015). Unlike above described abortive infection, this type of assembly interference still allows cellular processes to proceed, resulting in the formation of phage-like, but non-infectious particles (Tallent et al., 2007; Fillol-Salom et al., 2018). These SaPIs reside dormant in bacterial chromosomes, until they are induced

by the infection of specific helper phages and contribute to lateral gene transfer (Lindsay et al., 1998). Upon induction of PICIs, replication of target phage is limited by the elements of PICIs, which redirect phage replication to PICI replication and use phage encoded structural proteins to package PICI-specific DNA, resulting in the formation of non-infectious, phage-like particles (Ram et al., 2012). The PICIs have a well-conserved genetic organizations and are specially characterized by absence of structural and lysis genes (Fillol-Salom et al., 2018).

A similar PICI-like element (PLE) has been reported in Vibrio cholera as well. It has been shown that phage ICPI overcomes Vibrio PLE by encoding a CRISPR-Cas system in the phage genome, which targets the PLE genome (Seed et al., 2013).

Assembly of capsid-associated proteins

The Dar antirestriction proteins of P1 and IP1 antirestriction proteins of T4 are comparable in that both of these proteins are assembled into the mature virions and are injected into the host cells during the infection cycle to protect phage DNA from restriction (Iida et al., 1987; Rifat et al., 2008). Contrary to scarce information available on P1 head morphogenesis, there is wealth of information available on T4 head morphogenesis. There are several instances where phages are found to deliver proteins to host cells, in addition to the genetic material. The well-studied Salmonella enterica phage P22 also injects proteins following infection of the host cells (Jin et al., 2015).

The capsids of P22 contain three internal proteins (also called “pilot”, “ejection” or “E”

proteins”), which are essential for infection, and have been shown to be injected into the host cells (Jin et al., 2015). Both IPs of T4 and E proteins of P22 are assembled in the virion capsids during initial stages of capsid morphogenesis (Thomas & Prevelige, 1991;

Leiman et al., 2003).

The assembly pathway of prokaryotic viruses shares a significant similarity. The capsid morphogenesis in most bacteriophages and eukaryotic viruses such as herpesviruses and adenoviruses start with the formation of a protein shell that is later packed with the genetic material. The steps involved in capsid morphogenesis of different phages T4,

P22, HK97 are different, but in general this process requires portal, major capsid and scaffolding proteins. The portal proteins occupy one of the twelve vertices of icosahedral phage capsids and form a dodecameric ring (Aksyuk & Rossmann, 2011; Veesler &

Johnson, 2012). The portal proteins act as a nucleation factor and recruits major capsid and scaffolding proteins to form a protein procapsid shell of correct size and shape. The scaffolding protein core, which could be made of one or multiple proteins, determines the size and shape of the capsid for a phage. The scaffolding proteins are also required for the incorporation of internal proteins such as IPs of T4 and E proteins of P22 in the procapsid shell (Veesler & Johnson, 2012; Mateu, 2013). Since the morphogenesis pathway of P1 has not been studied in a great detail, there is no information on the role of portal and scaffolding proteins in capsid size determination and incorporation of other capsid proteins. However, by genetic and biochemical characterization, we have shown

that the virion-associated antirestriction components of P1 are assembled in the capsid following a definite pathway (Chapter II).

Questions to be addressed

Interest in P1 antirestriction system, DarB in particular, stems from the results of bioinformatic analysis that indicates that P1 DarB-like proteins are conserved (Gill et al.,

2011). Most of these DarB-like proteins are associated with mobile DNA elements such as plasmids, conjugative transposons, insertion sequences, integrative conjugative elements or genomic islands. This suggests that these DarB-like proteins might protect these DNA elements when they are mobilizing across hosts with different restriction background. In some cases, these DNA elements are associated with virulence factors, antibiotic resistance or phenotypic conversion, which raises public health concern, thus the genetic and biochemical study of DarB is of great significance (Gill et al., 2011).

Using P1 and its E. coli host with different type I R-M systems, the mechanism of how

P1 antirestriction system protects its genome from type I R-M system mediated DNA cleavage will be studied. As described above, phage P1 ejects Dar proteins into its host to protect its DNA from host type I R-M systems. The original study reported by Iida et al. in 1987 implies DarA and DarB as the constituent proteins of P1 antirestriction system. In addition, in 1998, Iida et al. reported that some genes in the darA operon affected capsid morphogenesis. However, the phage mutants used in those studies were generated using classic genetic tools, in which multiple genes could have been affected,

thus ascribing a phenotype to a particular gene was ambiguous. Throughout this study a

 Red-mediated genetic recombineering approach was used to construct isogenic gene deletions in P1 prophage and plasmid-based complementation system to verify the phenotypes. In Chapter II, several genes are interrogated to determine if components other than the darA and darB operons are important in the antirestriction system. It has been demonstrated that hdf, ddrA and ddrB from the darA operon and ulx from the darB operon form this multicomponent antirestriction system. Besides, it has been shown that these genes encode virion-associated proteins and these proteins are incorporated in the mature virions following a definite pathway. Moreover, the roles of hdf and darA in head-size determination of P1 virions have also been established. Building up from our findings described in Chapter II, P1 antirestriction system has been further characterized in Chapter III. Chapter III has three parts: In the first part, P1 genes of unknown functions have been tested for their role in protecting P1 DNA from type I R-M systems.

In the second part, it has been determined that the antirestriction proteins are incorporated in the virions before packaging of DNA. More importantly, a few N- terminal residues of the antirestriction protein DarB were found to be significant in directing the incorporation of DarB into P1 virions. In the third part, expression and purification of DarB have been optimized to obtain DarB for testing the biochemical mechanism of antirestriction function.

Understanding phage-host relationship is crucial for several reasons. Food and biotechnology industries use phage resistant bacterial strains to manufacture several

products. Thus, the emerging phage population, which could propagate on the selected bacterial strains, have to be constantly evaluated to ensure continuous product development (Labrie et al., 2010). Moreover, due to the emergence of antibiotic resistant bacteria, phage therapy, the use of phages to treat bacterial pathogens, is receiving renewed interest. Classical phage biology has relied on isolating novel phages from environmental sources for any bacterial strains. With the advances in systems and synthetic biology, the concept of synthesizing phages with a broader host range and host-independence have been proposed. As co-evolutionary arms race between phages and bacteria result in spectacular host-phage interactions, it is crucial to understand these relationships to successfully engineer any phages. Since bacteria can readily become resistant to phages by mutating the receptors, it is important to characterize the receptors so that phages targeting different receptors can selected for therapeutic purposes.

Because of the adaptability of phage genomes, most of the sequenced phage genes are of unknown function. Since the genetic material of phages is ejected into the host cells in each infection cycle, efforts should be made to understand the essentiality of these genes.

More importantly, phages also use internal host factors such as chaperones, transcription factors during infection cycle. These interactions must be understood especially in context of therapeutic phages so that robust phages that do not rely completely on host factors can be engineered and selection of phage resistant bacteria can be limited

(Labrie et al., 2010; Young & Gill, 2015).

Chapter IV discusses application of high-throughput genetic screen to study host-phage interactions of novel E. coli phages, LL5 and LL12, belonging to TLS- and rV5-like group, respectively. Receptors for these two phages have been characterized and it has been discovered that two chaperones, PpiB and SecB, are required for efficient infection cycle of phage LL5. In Chapter V, future directions of these research projects have been discussed.

CHAPTER II

THE MULTICOMPONENT ANTIRESTRICTION SYSTEM OF PHAGE P1 IS

LINKED TO CAPSID MORPHOGENESIS1

Introduction

Bacteriophage P1 was discovered by G. Bertani in 1951 as a temperate phage residing in

Escherichia coli strain “Li” (Bertani, 1951). P1 is a myophage of the order Caudovirales, with a 94 kilobase (kb) unit genome of linear dsDNA (Lobocka et al., 2004); unlike most other known temperate phages, P1 lysogenizes E. coli as a circular plasmid maintained at one copy per host chromosome (Ikeda & Tomizawa, 1968; Rosner, 1972). P1 has also played a major role in molecular genetics as the premier generalized transducing phage of E. coli (Lennox, 1955; Calendar, 1988).

In Caudovirales phages such as P1, the infection cycle begins with the adsorption of the phage to the host surface and the ejection of phage DNA into the host cell. Bacteria have several mechanisms to defend themselves against phage infection, including restriction and modification (R-M) systems that recognize and cleave foreign DNA in a site- specific manner (Fig. 2.1A). These systems are broadly divided into three categories, type I, II and III, which are distinguished by their subunit makeup, DNA cleavage

1 Reprinted with permission from “The multicomponent antirestriction system of phage P1 is linked to capsid morphogenesis” by Piya D., Vara L., Russell W.K., Young R. and Gill J.J., 2017. Mol Microbiol. 2017 August; 105(3): 399–412. Copyright by John Wiley and Sons, Inc. 37

mechanisms and the nature of their DNA recognition sequences (Kruger & Bickle, 1983;

Loenen et al., 2014).

Fig. 2.1. The antirestriction system of phage P1 protects DNA from type I restriction and modification (R-M) mediated cleavage. Reprinted with permission from Piya et al., 2017. Panel A. Phage infection begins with the translocation of the linear 95 kb P1 genome from the phage head into the host cytoplasm. Depending on the bacterial host where phages are propagated, specific bases in the phage DNA are methylated (denoted as stars). If the methylation pattern in the DNA matches the specificity of the host type I R-M system, the DNA is protected from cleavage (left), resulting in normal plating efficiency. If the phage DNA is unmethylated (or improperly methylated), the DNA is cleaved by the host type I R-M system (middle), resulting in reduced plating efficiency. Iida et al. (1987) demonstrated that in P1, virion-associated proteins (triangles) can protect P1 DNA from type I restriction, even if it is unmodified (right). It is believed that at least some of these virion-associated proteins must translocate into the cell in order to function.

Fig. 2.1 Continued. Panel B. Partial genomic map of P1 shows genes associated with the antirestriction system. Rectangular blocks represent genes, and those above the black line are transcribed rightwards, and genes below the line are transcribed leftwards. Besides the previously described darA and darB, several other genes are associated with the P1 antirestriction system. The dar genes (grey) are organized in two separate predicted transcriptional units. One transcript contains darB and ulx

(darB operon, driven by the PdarB promoter) and the other contains hdf, darA, ddrA and ddrB (darA operon, driven by Pdar). The ruler at the bottom of the figure corresponds to the nucleotide position as it appears in the published P1 genome (NC_005856.1).

Type I R-M systems are composed of three subunits, HsdR (R), HsdM (M) and HsdS (S)

(Hsd for host specificity of DNA). These subunits can form two different oligomeric complexes; R2M2S1, which can catalyze both restriction and modification, and M2S1, which can only catalyze modification (Suri et al., 1984; Murray, 2000). The HsdS subunit recognizes specific, bipartite DNA sequences of 13-15 bp, the HsdM subunit recognizes the modification status in the recognition DNA sequences and the HsdR subunit catalyzes DNA cleavage at a non-specific location up to a few kb away from the recognition sequence, if DNA is unmethylated or is methylated inappropriately (Murray,

2000). Unlike type I R-M systems, type II R-M systems have independent restriction and modification enzymes. These restriction enzymes cleave DNA basepairs within the recognition sites if the DNA is not appropriately modified (Bickle & Kruger, 1993).

Similar to type II R-M systems, type III systems are comprised of separate modification and restriction subunits. The modification subunit can function by itself as a modification methylase. However, a complex of modification and restriction subunits is required for endonuclease function, in which DNA cleavage occurs 25-27 bp away from the recognition site (Tock & Dryden, 2005).

P1 encodes the virion-associated proteins DarA and DarB (dar standing for defense against restriction), which are required to protect the P1 genomic DNA (gDNA) from restriction by host type I R-M systems (Fig. 2.1A) (Iida et al., 1987). DarA, with a predicted molecular mass of 69 kDa, is expressed late in the lytic cycle and is processed into a smaller 60 kDa form when incorporated into the virion (Streiff et al., 1987). DarB

has a predicted molecular mass of 251 kDa and appears to be incorporated into the virion intact (Iida et al., 1987). Iida et al. demonstrated by plaque assay that DarA is required for protection of DNA against restriction by the EcoA type I R-M system and DarB is required for protection against the EcoB and EcoK systems. The efficiency of plating

(EOP) of P1 darA mutants was reduced in EcoA, EcoB and EcoK strains whereas the

EOP of P1 darB mutants was reduced only in EcoB and EcoK strains. Because complementation studies indicated that the Dar proteins function only in cis, it has been proposed that these are ejected into the host cell along with phage DNA at the initiation of infection, where they exert their antirestriction activity (Fig. 2.1A) (Iida et al., 1987).

The antirestriction proteins synthesized during the phage lytic cycle only act in the following infection cycle, and the antirestriction activity is not specific to P1 gDNA, but any DNA that is packaged into the P1 virion (Iida et al., 1987). Since darA- phages also exhibit the darB- phenotype and darA- virions lack both DarA and DarB proteins, it appears that DarA is required for the incorporation of DarB into the P1 capsid (Iida et al., 1987). The 2,255-residue DarB contains an identifiable N-terminal methyltransferase domain and a central DExD-like helicase domain, enzymatic functions that could be imagined to play a direct role in the protection of DNA from restriction (Lobocka et al.,

2004). Bioinformatic analysis of DarB revealed that it is widely distributed among bacteria, primarily associated with mobile DNA elements including phages, plasmids and conjugative transposons (Gill et al., 2011). It has been proposed that DarB could represent a larger class of proteins that facilitate DNA mobility by protecting mobile

DNA elements from type I R-M system-mediated cleavage during their transfer between hosts.

To further explore the antirestriction system of phage P1, we used a recombineering approach to create isogenic deletion mutants of darA, darB, and other P1 genes with previously unknown function including hdf, ddrA, ulx and ddrB. The results are discussed in terms of a network of proteins involved in the assembly of P1 antirestriction system and their effects on capsid morphogenesis.

Materials and methods

Bacterial strains and phages

The bacterial strains and the parent phage P1CMclr100 used in this study were obtained from the Coli Genetic Stock Center, Yale University, and are listed in Table 2.1. P1CM is a derivative of P1kc that acquired chloramphenicol resistance from the R-factor R14

(Kondo & Mitsuhashi, 1964). P1CMclr100, hereafter referred to simply as P1, is a thermoinducible mutant of P1CM (Rosner, 1972). All phage mutants used in this study are single-gene deletions of P1. Unless otherwise noted, E. coli strains were cultured on

LB broth [10 g L-1 Bacto Tryptone (BD), 5 g L-1 Bacto yeast extract (BD), 10 g L-1 NaCl

(Avantor)] or LB agar (LB broth amended with 15 g L-1 Bacto agar) at 37 °C. P1 lysogens were cultured and maintained at 30 °C on LB amended with 10 µg mL-1 chloramphenicol (LB cm) or 10 µg mL-1 chloramphenicol plus 30 µg mL-1 kanamycin

(LB cm+kan).

Table 2.1. Bacterial strains, phages and plasmids. Reprinted with permission from Piya et al., 2017. Strains, phages or Genotype or relevant characteristic Reference/source plasmids E. coli strains - - - - - WA2379 leu met lac rA mA (Arber & Wauters-Willems, 1970) / The Coli Genetic Stock Center (CGSC), Yale University - - + + W3110 F  rpoS(Am) rph-1 Inv(rrnD-rrnE) rK mK (Iida et al., 1987; Hayashi et al., 2006) / CGSC ------WA921 thr leu met lac rK mK (Wood, 1966; Arber & Wauters-Willems, 1970) / CGSC + - - - + + WA960 thrB leu met lac rB mB (Wood, 1966; Arber & Wauters-Willems, 1970) / CGSC BW25113(pKD46) F- l- rpoS(Am) rph-1 rrnB3 ΔlacZ4787 (Baba et al., 2006) / CGSC hsdR514 Δ(araBAD)567 Δ(rhaBAD)568 q BW25141(pKD4) lacI rrnBT14 ΔlacZWJ16 ΔphoBR580 hsdR514 (Datsenko & Wanner, 2000) / ΔaraBADAH33 ΔrhaBADLD78 galU95 endABT333 CGSC uidA(ΔMluI)::pir+ recA1

MG1655 lacIq1tonA::Tn10 (Park et al., 2006) / Lab stock Phages P1ΔdarB in-frame deletion of darB in P1CMclr100 This study P1Δulx in-frame deletion of ulx in P1CMclr100 This study P1ΔdarA in-frame deletion of darA in P1CMclr100 This study P1Δhdf in-frame deletion of hdf in P1CMclr100 This study P1ΔddrA in-frame deletion of ddrA in P1CMclr100 This study P1ΔddrB in-frame deletion of ddrB in P1CMclr100 This study P1Δhxr in-frame deletion of hxr in P1CMclr100 This study Plasmids pdarB P1 darB cloned into pBAD24 This study pulx P1 ulx cloned into pBAD24 This study pdarA P1 darA cloned into pBAD24 This study phdfam_darA_ddrAam P1 hdf_darA_ddrAgene fragment cloned into This study pBAD24 and amber mutations introduced to 9th codon in hdf and 28th codon in ddrA phdf P1 hdf cloned into pBAD24 This study pddrA P1 ddrA cloned into pBAD24 This study pddrB P1 ddrB cloned into pBAD24 This study pulx-CFLAG P1 ulx with C-terminal FLAG-tag fusion This study cloned into pBAD24 43

Production of phage lysates

Phage lysates were produced by thermal induction of P1 lysogens. Lysogenic strains were grown at 30 °C in LB cm to OD550 0.5 - 0.6. P1 was thermally induced by shifting the culture to 42 °C in a shaking water bath for 60-75 min (Iida & Arber, 1977). The crude lysate was harvested when the OD550 fell to ~0.2, by centrifugation of the culture at 10,000 x g, 10 min, 4°C and sterilized by passage through a 0.22 µm syringe filter

(Millipore).

Lysogenization

Lysogens of P1 and its mutants were produced as previously described (Rosner, 1972) with minor modifications. A fresh overnight culture of the desired E. coli lysogenization host was supplemented with 5mM CaCl2, and 100 µL of the culture was mixed with 100

µL of an undiluted phage lysate (~108-109 PFU mL-1) and incubated at RT for ~20 minutes. The phage-host mixture was then plated to LB cm and a CMR colony was selected after overnight incubation at 30 °C and purified by restreaking. The same procedure was used for producing lysogens of P1 deletion mutants, with the exception that plating was conducted on LB cm+kan.

Generation of PCR fragments for single-gene deletions

All primers (table 2.2) used in PCR were ordered from Integrated DNA Technologies.

The plasmid pKD4 has an FRT-flanked kan gene which was used at the source of the kanamycin resistance cassette used for gene deletions (Datsenko & Wanner, 2000). To 44

Table 2.2. Primers. Reprinted with permission from Piya et al., 2017. Forward primer 5'-3' Reverse primer 5'-3' Usage tttgtcaaAaccgacctgtccg ggacaggtcggtTttgacaaaa To mutagenize EcoP1 site in ag kanamycin resistance gene in pKD4. The nucleotides in capital letters represent the mutation that is introduced. ggttttatggacagcaagcg gcttccatccgagtacgtg Sequencing primers to verify site-directed mutation in pKD4 tggcgaactcaaacgtcgtctta gaggtccattaactcaatagct To knockout darA from P1 agcaactgaaagcaggaaattaa gatagtgtcatgctgctacccc genome catggtgtaggctggagctgctt cgctttatgggaattagccatg c gtcc gtctttcaatcaacagcagatca acaccatcatgttccgaagg Sequencing primers to verify c darA deletion in P1 genome tcgtgtttgaacggaatttaaca tgcgacgcccggcgaaccgggc To knockout darB from P1 ctagtcacttgttaaggattacc gctcctgttatgcgtattgttg genome aatggtgtaggctggagctgctt gatgacatgggaattagccatg c gtcc aggaggatgttgtcccgttc tttcagtaatcgcccgcgtag Sequencing primers to verify darB deletion in P1 genome gcagttacgtaataaatctcgca atccctttattgatattgaact To knockout hdf from P1 caggatgtgtcagatgacgaaaa gttccatgttaatttcctgctt genome ataagtgtaggctggagctgctt tcagttatgggaattagccatg c gtcc tatatgggtaaaggagagcgact tccgtagcacaccagtggatc Sequencing primers to verify g hdf deletion in P1 genome acctggctgatctgctggtagca tgatcgcttaagctcatccgtg To knockout ddrA from P1 atccagaaagcgggggtagcagc ttacgccttatgctgcctcttt genome atgagtgtaggctggagctgctt aatgttatgggaattagccatg c gtcc ggatatgcaaagcactgacatgg ccgtccagagtattggatacc Sequencing primers to verify ddrA deletion in P1 genome tgctggaactgttattaggtgtg gaacaggttatttttgcactat To knockout hxr from P1 gcaaaggagctaaataatgcaga caggtagttaatatactatctg genome ttaagtgtaggctggagctgctt gtctacatgggaattagccatg c gtcc tatacccgccagcctcagtaag ctgcggcggttataggttcc Sequencing primers to verify hxr deletion in P1 genome cccggttcgccgggcgtcgcata cccatgctttactgtgtagtat To knockout ulx from P1 atatggccacactatctgataca cgctttttcaagttttctctcc genome ataagtgtaggctggagctgctt agcctgatgggaattagccatg c gtcc

Table 2.2. Primers. Continued Forward primer 5'-3' Reverse primer 5'-3' Usage cgaaagacgcaatcaagacg tgatctgctcccattcttcg Sequencing primers to verify ulx deletion in P1 genome catggatagccgccaacattaaa atctccgcctaatggcgctgcg To knockout ddrB from P1 gaggcagcataaggcgtaacacg attttaatctgcattatttagc genome gatggtgtaggctggagctgctt tcctttatgggaattagccatg c gtcc agtggcgaggagaaacaacc tcaagggtgtcaacgaatcc Sequencing primers to verify ddrB deletion atcgtctagagtaataaaggagg atcgaagcttttatgcgtattg Clone darB into pBAD24 tatcgatatgaacaagctatcta ttggatgacggc tgggggtg atcgatctagagtaaaggagatc atcgaaagcttttatgctgcct Clone hdf, darA and ddrA gatcatgtgtcagatgacgaaaa ctttaatgttggc into pBAD24 ataagtatgc atcgatctagagtaaaggagatc acacaaagcttttaatttcctg Clone hdf into pBAD24 gatcatgtgtcagatgacgaaaa ctttcagttgcttaag ataagtatgc atcgatctagagtaaaggagatc atcgaaagcttagctgatagtg Clone darA into pBAD24 gatcatggaacagttcaatatca tcatgctgc ataaaggg atcgatctagagtaaaggagatc atcgaaagcttttatgctgcct Clone ddrA into pBAD24 gatcatgacactatcagctattg ctttaatgttggc agttaatgg atcgatctagagtaaaggagatc atcgaaagctttcaagttttct Clone ulx into pBAD24 gatcatggccacactatctgata ctccagcctgtg caataaaacc atcgatctagagtaaaggagatc atcgaaagcttttaatctgcat Clone ddrB into pBAD24 gatcatgagcttaagcgatcagg tatttagctcctttgcc tgg gaaaaataagtaGgcaacggtcg tcgaccgttgcctacttatttt To make amber mutation in attttg tcg the 9th codon of hdf cgagtggctaggagttgctg caactcctagccactcgcgg To make amber mutation in the 28th codon of ddrA atcgatctagagtaaaggagatc atcgaaagctttcacttgtcgt To clone ulx with C-terminal gatcatggccacactatctgata catcgtctttgtagtcagaacc FLAG-tag fusion into caataaaacc agttttctctccagcctgtgc pBAD24

avoid restriction by the Type III R-M system resident in the P1 chromosome, an EcoP1 site normally present in the kan gene was removed by following protocol from

QuikChange site-directed mutagenesis kit (Agilent Technologies) changing G608 to A in a silent mutation. To generate FRT-flanked kan insert for gene deletion, the 3’-ends of the forward and reverse primers were designed to match the priming sites gtgtaggctggagctgcttc and atgggaattagccatggtcc of pKD4 (Datsenko & Wanner, 2000).

The 5’ ends of both forward and reverse primers were synthesized to include 50 nt homology to the P1 genome up- and downstream of the targeted gene. The flanking regions were chosen to retain the first and last several codons of the targeted genes to avoid polar effects (Datsenko & Wanner, 2000). PCR reactions were performed using

Phusion Hi-Fidelity PCR Master Mix (New England Biolabs), following the manufacturer’s recommended protocol. The PCR products were gel purified by using the

QIAquick Gel Extraction Kit (Qiagen).

Generation of single-gene knockout mutants

The phage lambda Red recombinase mediated homologous recombination method was used to generate isogenic single-gene knock-out P1 mutants (Datsenko & Wanner,

2000). P1 was lysogenized into BW25113(pKD46) and a colony resistant to both chloramphenicol and ampicillin was selected. BW25113(pKD46) lysogenized with P1 was grown in LB amended with 100 µg mL-1 ampicillin and 10 µg mL-1 chloramphenicol to OD550 0.1, 1 mM L-arabinose was added to induce Red proteins, and the culture was grown to OD550 0.6. The cells were harvested and made

electrocompetent as described previously (Datsenko & Wanner, 2000). Each electroporation reaction contained 300 ng of gel-purified DNA and 100uL competent cells in a 0.1 cm cuvette and was transformed in a Bio-Rad MicroPulserTM electroporator following the manufacturer’s protocol. Cells were recovered in 1 mL SOC for 2 hours at

30 oC and plated to LB cm+kan at 30 °C overnight. CMR and KanR colonies were selected and mutations were verified by PCR amplifying a DNA region spanning the kan insertions and sequencing across the insertion junctions (Baba et al., 2006). These lysogens were then induced as above and the mutant P1 phages were lysogenized into E. coli strain MG1655 as described above for maintenance of the strains.

Complementation of phage mutants

Phage knockouts were complemented in trans by induction of lysogens containing both the mutant prophages and vectors expressing the deleted genes. Complementing genes were amplified by PCR from a P1 DNA template (Table 2.1) and cloned into pBAD24 at its XbaI and HindIII sites using standard molecular biology techniques (Guzman et al.,

1995). The primers used for PCR are listed in table 2.2. Ligation products were transformed into competent E. coli 5-alpha cells (New England Biolabs) and selected by plating on LB agar amended with 100 µg mL-1 ampicillin. The plasmids were extracted as described above and were verified by sequencing before transformation into the r-m- strain WA921. Complemented phages were prepared by thermal induction of mutant phage from WA921 lysogens containing the corresponding complementing vector expressing the deleted gene in trans (Table 2.1). 1 mM L-arabinose (Sigma-Aldrich) was

added to the culture at the time of temperature shift to 42 °C to induce protein expression from the complementing plasmids. The phage lysate was used to determine Efficiency of

Plating (EOP) on restricting and non-restricting hosts as described below.

Efficiency of Plating (EOP) Assay

All phages used in EOP assays were induced from modification-deficient WA921 lysogens. Assays were conducted as previously described with few modifications (Arber

& Dussoix, 1962; Mise & Arber, 1976; Iida et al., 1987). Host cells, grown to OD550 0.4

- 0.5 in tryptone broth (10 g L-1 Bacto tryptone, 5 g L-1 NaCl) amended with appropriate antibiotics, were incubated with 10 mM CaCl2 for 30 minutes at RT. Phage were adsorbed to 300 μL of host cells for 20 minutes at RT. The cells and phage were then plated using the soft agar overlay method using LB (Lennox) plates (10 g L-1 Bacto tryptone, 5 g L-1 NaCl, 5 g L-1 Bacto yeast extract, g L-1 Bacto agar) containing 2.5 mM

-1 CaCl2 as the bottom plates with 4 mL lawns of tryptone top agar (10 g L Bacto tryptone, 5 g L-1 NaCl, 7 g L-1 Bacto agar). Plaques were counted after overnight incubation at 42 °C. EOP was calculated as the ratio of plaques appearing on the lawn of the restricting strain to the number of plaques on WA921 lawns. The EOP of each phage mutant was normalized to the EOP of parental P1 on the same plating strain. In complementation assays, EOPs were normalized to P1 induced from WA921(pBAD24).

All experiments were replicated three times.

Purification of virions by CsCl isopycnic centrifugation

P1 or P1 mutant was induced in 1L LB as described above. The crude lysate was centrifuged in JA10 rotor for 15 minutes at 17000 x g and the supernatant was filter sterilized. The lysate was concentrated by 24-hour centrifugation in JA10 rotor at 14000 x g at 4°C. The pellet was soaked in SM buffer (50 mM Tris-HCl pH 7.5, 100 mM NaCl,

-1 8 mM MgSO4) at 4°C and extracted after 48 hours. 10 µg mL DNase (Sigma) was added to the concentrated phage and left at RT for 30 minutes. Phages were then purified by equilibrium centrifugation in Cesium chloride as previously described (Boulanger,

2009). The phage bands were extracted with 18-gauge needles and dialyzed against SM buffer in Slide-A-Lyzer 3500 MWCO dialysis cassettes (Thermo Scientific). Small volumes of pre-dialysis samples were saved and used to measure refractive index in an

Abbe Refractometer. Refractive indices were converted to density (g cc-1) by comparing to standard CsCl density-refractive index correlation table.

SDS-PAGE analysis

For SDS-PAGE analysis of phage proteins, samples were prepared as described previously, with slight modification (Boulanger, 2009). Approximately 2 x 1010 PFU of

CsCl purified P1 were loaded per lane. For antirestriction P1 mutants, protein loading was normalized to equal amounts of tail sheath protein (57 kDa). Phages were heated in boiling water for 10 minutes to release DNA from the capsid and samples were then treated with DNase I at 37 °C for 2 hours. The samples were denatured by heating in boiling water for 5 minutes in Laemmli sample buffer and loaded on a 4-20% Tris-

glycine SDS-PAGE gel (Life Technologies) (Laemmli, 1970). PageRuler Unstained

Broad Range Protein Ladder (Thermo Scientific) was used as molecular mass standard.

The gel was stained with SYPRO Ruby (Thermo Scientific), following the manufacturer’s recommended protocol for maximum sensitivity. The gel was imaged with Fotodyne gel imager.

Proteomic analysis

Proteins associated with the virion were identified by mass spectrometry (MS) of the protein bands excised from SDS-PAGE gel as described previously (Gill et al., 2011).

Briefly, ~1011 PFU of phage P1 was prepared and loaded into 4-20% Tris-glycine SDS-

PAGE as described above. Coomassie-stained bands were excised, subjected to reduction with dithiothreitol and alkylation with iodoacetamide. The samples were treated with ~0.4 µg trypsin (Thermo Scientific) and digested peptides were concentrated and desalted using C18 ZipTips (Millipore). The samples were spotted manually onto a matrix-assisted desorption ionization (MALDI) target (Genomic

Solutions) using α-cyano 4-hydroxycinnamic acid. MALDI-MS and MS/MS analysis were conducted against the NCBI-nr database as described previously (Gill et al., 2011).

P1 DarB, DarA and Hdf were identified by this method.

The virion-associated DdrB was identified by liquid chromatography-tandem mass spectrometry (LC-MS/MS) in collaboration with the Protein Chemistry Lab at Texas

A&M University. Four lanes each of ~2 x 1010 PFU P1 and P1ΔddrB were loaded into a

4-20% Tris-glycine SDS-PAGE and stained with SyproRuby as described above. The gel region missing protein band in P1ΔddrB lanes and corresponding protein band from

P1 lanes were excised. The samples were processed for LC-MS/MS as described before

(Shevchenko et al., 2006). LC-MS/MS was performed on a LTQ Orbitrap Velos/ETD mass spectrometer (Thermo Fisher) equipped with a NanoLC 2-D HPLC system

(Eksigent). The results were analyzed using Mascot (Matrix Science) and X!Tandem

(The GPM). Scaffold (Proteome Software) was used to validate MS/MS based peptide and protein identifications. The SDS-PAGE band was annotated based upon its presence in P1 and absence in P1ΔddrB lanes.

Transmission Electron Microscopy

Phages were stained with 2% uranyl acetate and imaged in a JEOL 1200 EX transmission microscope under 100 kV accelerating voltage as previously described

(Valentine et al., 1968; Gill et al., 2011). Side-to-side head diameters perpendicular to the axis of the tail were measured electronically using ImageJ (Schneider et al., 2012) and converted to nm against images of a carbon grating replica of known dimensions

(Ted Pella, cat# 607).

Results and discussion

Genomic context of the P1 antirestriction system

The genome of phage P1 was completed in 2004 (NC_005856.1), showing the positions of the darA and darB structural genes within the P1 chromosome (Lobocka et al., 2004).

As shown in the partial P1 genetic map (Fig. 2.1B) the 1,920 bp darA gene is located in a polycistronic operon with its expression driven by LPdar (LP for late promoter)

(Guidolin et al., 1989b), and darB (6,768 bp) is located ~10 kb downstream of darA in a separate operon driven by PdarB (Lobocka et al., 2004). The darA gene is located immediately downstream of hdf, and upstream of ddrA, ddrB and hxr. The genes lydA and lydB, upstream of hdf, are known to function as the phage holin and antiholin, respectively (Schmidt et al., 1996). Two operons are located between the dar operons: one containing the transcript of the phage’s SAR endolysin lyz (Xu et al., 2004) and another on the opposite strand encoding the predicted phage portal, portal protease and single-stranded DNA binding protein (Lobocka et al., 2004). The darB gene is upstream of ulx and the lysogeny maintenance gene lxc (Lobocka et al., 2004).

The genes shown in grey in Fig. 2.1B were deleted and replaced with kan markers by a lambda Red recombineering approach (Datsenko & Wanner, 2000). Initially darA and darB were deleted, followed by hdf, ddrA, ddrB, hxr and ulx. Because lydAB, lyz, lxc and the phage structural proteins Prt and Pro already had assigned functions, they were not considered to be relevant to this study and were not studied further. Deletion of hxr produced no detectable antirestriction phenotype against EcoA, EcoB or EcoK (data not shown) and genes downstream of hxr were not examined further.

darB and ulx provide protection against EcoB and EcoK

The antirestriction phenotypes associated with P1 darA and darB were first reported in

1987 (Iida et al., 1987): P1, when propagated on a modification-deficient host, exhibited a slight (less than ten-fold) reduced efficiency of plating (EOP) on EcoA and EcoK lawns and an EOP of ~10-2 on EcoB. Under the same conditions, P1darA- plated at an

EOP ~10-4 to 10-5 on all three restrictive hosts, whereas P1darB- was restricted (EOP ~

10-4) only on EcoB and EcoK.

The previously reported darB- phenotype, defined as severely reduced plating efficiency on EcoB and EcoK hosts, was recapitulated in an isogenic deletion of darB (Fig. 2.2A).

As previously reported, P1∆darB did not exhibit an antirestriction defect in the EcoA host. In addition to reproducing the darB phenotype, the antirestriction defect in

P1∆darB could be efficiently complemented in trans by inducing the mutant phage from a lysogen of the non-modifying host strain WA921 carrying the plasmid pdarB, expressing DarB from an arabinose-inducible promoter (Fig. 2.2A). In this complementation system, the progeny phage could incorporate the DarB protein expressed from the plasmid in the propagating host and produce the antirestriction phenotype upon infection of a restricting strain. Phages complemented in this way could only express the antirestriction phenotype for their first infection cycle. Complemented mutant phage that were re-propagated on WA921 exhibited the same EOP as the original

P1∆darB when plated to all three of the restricting strains (data not shown), indicating the absence of recombination between the phage and complementing vector during

A B

Fig. 2.2. darB and ulx are required for protection of P1 against EcoB and EcoK restriction. Reprinted with permission from Piya et al., 2017. Parental P1 or isogenic deletion mutant phages were thermally induced from lysogenized modification-deficient WA921 containing either empty pBAD24 or the respective complementing plasmid. Phages were then plated on E. coli hosts containing the Type I EcoA, EcoB, or EcoK R- M systems and WA921; efficiency of plating (EOP) for each phage was calculated as the ratio of plaques formed on lawns of the R-M-containing strains to the plaques formed on WA921. The EOP data shown have been normalized to the EOP of parental P1 induced from WA921 containing pBAD24. Panel A: the relative EOP of P1ΔdarB -3 -4 was reduced by ~10 and 10 in EcoB and EcoK strains, respectively (gray bars). -2 -3 Panel B: the relative EOP of P1Δulx was reduced by ~10 and 10 in EcoB and EcoK strains respectively (gray bars). The restriction phenotype associated with both darB and ulx could be complemented in trans (white bars). No notable phenotype was observed in the EcoA strain. The data shown are averages of three biological replicates and error bars represent standard deviation.

phage propagation. The reduced EOP of P1∆darB in EcoB and EcoK cells indicates that in the absence of DarB, P1 DNA, after being ejected into host cells, is susceptible to

EcoB and EcoK restriction, but is protected from EcoA restriction.

Other genes near darB were examined for their possible role in antirestriction. The gene ulx (146 codons; ulx for upstream of lxc (Lobocka et al., 2004)) is located between darB and lxc and was previously of unknown function. P1∆ulx showed a partial darB-like restriction phenotype, with EOP attenuated approximately 10-fold less than for P1∆darB

(Fig. 2.2B). The ulx-associated restriction phenotype could also be complemented in trans (Fig. 2.2B). In both darB and ulx deletions, complementation produced slightly higher EOP’s than the parent P1 phage, suggesting complementation could provide a greater copy number of protein available for incorporation into the virion and subsequent greater protection against restriction. The plaque sizes for both P1ΔdarB and P1Δulx were comparable to P1. Moreover, a double deletion mutant of darB and ulx showed plating deficiency similar to P1ΔdarB (data not shown). Thus, both darB and ulx are required for full protection of P1 DNA against EcoB and EcoK restriction.

darA, hdf and ddrA contribute to protection against EcoA, EcoB and EcoK

In agreement with previous work, the isogenic deletion P1∆darA had reduced EOP when plated on strains expressing EcoA, EcoB or EcoK (Fig. 2.3A). Relative to P1, the EOP of P1∆darA was decreased by ~10-3, 10-2 or 10-4 in cells expressing EcoA, EcoB or

EcoK, respectively. The EcoA-associated darA restriction phenotype could be only

A B C

Fig. 2.3. hdf, darA and ddrA are required for protection of P1 against EcoA, EcoB and EcoK restriction. Reprinted with permission from Piya et al., 2017. Phages were induced and EOP assays were performed as described previously. The EOP data shown have been normalized to EOP of parental P1 induced from modification deficient WA921 containing pBAD24. Panel A: The relative EOP of P1ΔdarA was reduced by -3 -2 -4 ~10 , 10 and 10 in EcoA, EcoB and EcoK strains respectively (dark gray bars). The darA-associated restriction phenotype could be complemented partially by a plasmid bearing only darA (light gray); stronger complementation was observed with a plasmid expressing a longer transcript containing darA, along with the upstream gene hdf , and am downstream ddrAam (white bars). Panels B and C: Both P1Δhdf and P1ΔddrA exhibited a darA-like restriction phenotype (gray bars). The restriction phenotypes of both hdf and ddrA could be complemented in trans (white bars). The data shown are averages of three biological repeats and the error bars represent standard deviation.

partially complemented in trans by darA alone, and the EcoB and EcoK phenotypes were not affected. It was previously reported that darA mutants could be complemented to nearly wt levels in trans by a P1 DNA fragment containing darA, along with the upstream gene hdf and the downstream gene ddrA of the darA operon (Iida et al., 1998).

In our experiments, P1∆darA was more efficiently complemented in trans by a construct

containing darA, hdf and ddrA (data not shown). Better complementation in presence of hdf and ddrA raised a possibility of the polar effects of darA deletion, in which the restriction phenotype observed could either be associated with hdf or ddrA, the other genes present in the complementing plasmid. In order to rule out the roles of Hdf and

DdrA in complementation, nonsense mutations were introduced in both hdf and ddrA to prevent their expression. Same degree of complementation was observed with phdfam_darA_ddrAam (Fig. 2.3A). The low efficiency of complementation exhibited by the pdarA vector may be due to instability of the mRNA transcript containing darA alone, as the longer DNA fragments included on phdfam_darA_ddrAam and the previously reported P1 DNA fragment provided better complementation.

Previous studies of darA and darB function used P1 mutants that contained either IS insertions or deletions that often spanned hdf and ddrA (Iida et al., 1987). To determine if these genes could also play a role in the protection of P1 DNA from host restriction, the isogenic mutants P1∆hdf and P1∆ddrA were created and evaluated for their antirestriction phenotypes. Both P1∆hdf and P1∆ddrA showed darA-like phenotypes, with sensitivity to EcoA, EcoB and EcoK restriction similar to that exhibited by

P1∆darA (Fig. 2.3B and 2.3C). Unlike darA, the hdf and ddrA phenotypes could both be fully complemented in trans by vectors containing hdf or ddrA alone (Fig. 2.3B and

2.3C). The plaque sizes for both P1∆hdf and P1∆darA were smaller than that of P1, whereas the plaques of P1∆ddrA appeared to be of normal size.

ddrB modulates the EcoB and EcoK antirestriction phenotype

Having determined the restriction phenotypes in ulx, hdf and ddrA mutants, we tested other genes located within the darA and darB operons (Fig. 2.1B). Surprisingly an isogenic deletion of ddrB was found to have a ~10-fold higher EOP on cells expressing

EcoB or EcoK than that of the parental P1 (Fig. 2.4). This phenotype could again be complemented in trans by a vector containing ddrB alone, with the EOP returning to ~3 in the complemented phage. Deletion and complementation of ddrB produced no notable phenotype on EcoA strains. Both ulx and darB are similar to ddrB in this regard, in that their deletion affects susceptibility to EcoB and EcoK restriction, with no apparent effect on EcoA strains (Fig. 2.2). It seems likely that ddrB exerts its phenotype via action on

Ulx and/or DarB, either by regulating the copy number of these proteins incorporated into the virion, or by modulating their activity after the antirestriction system is deployed in the host cell.

Virion proteomics of P1 and its mutants

It was shown previously that P1darA- phages fail to package DarB (Iida et al., 1987), leading to the conclusion that, in addition to being required for protection of DNA against EcoA, DarA is also required for incorporation of DarB into the virion. In the present study, multiple genes in addition to darA and darB have been identified with roles in the antirestriction phenotype of P1. These phenotypes can be placed into two broad categories: darA-like, which affects phage EOP on EcoA, EcoB and EcoK strains,

and darB-like, in which EOP is affected only on EcoB and EcoK strains. In order to determine how these multiple P1 genes could produce similar restriction phenotypes,

Fig. 2.4. ddrB negatively affects the darB phenotype. Reprinted with permission from Piya et al., 2017. Phages were induced and EOP assays were performed as described previously. The EOP data shown have been normalized to EOP of parental P1 induced from modification-deficient WA921 containing pBAD24. Unlike the other restriction phenotypes described, disruption of ddrB increased the relative EOP of the phage in EcoB and EcoK strains by approximately 10-fold compared to wild-type P1 (gray bars). This effect did not extend to EcoA strains, suggesting that DdrB negatively impacts the activity of DarB. The ddrB phenotype could also be complemented in trans (white bars).

proteomic analysis of purified virions was conducted (Fig. 2.5). Protein bands corresponding to DarB, Hdf, DarA and DdrB could be assigned by mass spectrometry analysis (underlined in Fig. 2.5A). The protein band corresponding to DdrA is assigned based on missing band density corresponding to its predicted protein molecular mass in lanes of P1ΔddrA virions compared to the parental P1. The protein band corresponding to Ulx was assigned based on western blotting of purified P1Δulx virions complemented with a Ulx-FLAG fusion (Inclan et al., 2016) (Fig. 2.6).

Disruption of darB resulted in the loss of DarB and Ulx (Fig. 2.5), indicating only Ulx is dependent on DarB for incorporation. Coupled with the conserved domains present in

DarB, the EcoB and EcoK phenotypes observed for P1∆darB (Fig. 2.2A) suggests that

DarB is directly responsible for EcoB and EcoK antirestriction activity. Band densitometry indicates P1∆ulx packages only ~15% of the DarB as the parental P1 (data not shown). Thus, DarB can be packaged into the virion in the absence of Ulx but packaging efficiency is greatly reduced. This reduced packaging of DarB is consistent with the ~10-fold weakened antirestriction phenotype exhibited by P1∆ulx compared to

P1∆darB (Fig. 2.2).

Fig. 2.5. Comparative SDS-PAGE of parental P1 and isogenic mutants suggests cascading dependencies for protein incorporation into the P1 virion. Reprinted with permission from Piya et al., 2017. Panel A: SDS-PAGE of parental P1 and antirestriction mutants. Molecular masses of the size standard are provided in kDa on the left. Relevant protein bands are labeled on the left; proteins that have been identified by mass spectrometry are underlined. DdrA is assigned based on the missing band density in the gel image and predicted molecular mass, and Ulx is assigned based on western blotting of P1Δulx complemented with Ulx-FLAG with anti-FLAG antibody. At the bottom of the figure, the restriction sensitivity phenotypes of the P1 mutants are indicated: B, K (sensitive to EcoB and EcoK restriction), or A, B, K (sensitive to EcoA, EcoB and EcoK restriction).

B P1 P1 and antirestriction mutants proteins P1 ∆darB ∆ulx ∆ddrB ∆hdf ∆darA ∆ddrA DarB + - (+) + - - - DarA + + + + - - + DdrB + + + - - - - Hdf + + + + - - + Ulx + - - + - - - DdrA + + + + - - -

Fig. 2.5 Continued. Panel B: The table summarizes protein presence or absence in P1 mutants. P1Δulx shows a faint DarB band, which suggests that DarB can be incorporated in the absence of Ulx, albeit inefficiently. When either hdf or darA is disrupted, P1 fails to package both Hdf and DarA, suggesting that Hdf and DarA are co-dependent for virion incorporation.

20kDa 15kDa 10kDa

Fig. 2.6. Identification of Ulx band in SDS-PAGE of P1 virions. Reprinted with permission from Piya et al., 2017. Ulx protein band in P1 virions was identified by western blot on P1∆ulx complemented with functional Ulx fused to FLAG-tag on C- terminal end. The protein samples for SDS-PAGE were normalized and prepared as described in materials and methods. The samples were loaded in 4-20% Tris-glycine

(Thermo Scientific) in order as shown above. After SDS-PAGE, the gel was cut into two halves. One half was stained with SyproRuby as described earlier and the other half was used for western blot following standard protocol (Green & Sambrook, 2012). In brief, SDS-PAGE resolved proteins were transferred to PVDF membrane (GE

Healthcare) overnight at 30V. The membrane was blocked with 1% blocking buffer and probed with Anti-FLAG primary antibody (Thermo Scientific) and HRP-conjugated secondary antibody. The membrane was developed with SuperSignal West Femto Maximum Sensitivity Substrate (Thermo Scientific) and imaged (BioRad).

Compared to the parental P1, a single ~50 kDa band is missing in the P1∆ddrB virion; this species was confirmed as DdrB by mass spectroscopy analysis. Aside from this 50 kDa species, all bands including those corresponding to DarB, Ulx, DarA, Hdf, and

DdrA are present, indicating that no other virion proteins are dependent upon DdrB for their incorporation. Since ddrB encodes a protein with a predicted molecular mass of

~109 kDa, the appearance of DdrB at a position corresponding to 50 kDa suggests that the protein is proteolytically processed for incorporation into the virion; no ~109 kDa band corresponding to DdrB was observed in SDS-PAGE. Based on MS-MS analysis of the gel slice containing this predicted DdrB band from the parental P1, high peptide coverage (5 to 27-fold, covering 88% of residues) was observed for the N-terminal portion of the protein sequence, with coverage dropping abruptly after residue 421 (1 to

3-fold, covering ~42% of residues), suggesting that it is primarily the N-terminal portion of the DdrB protein that is present in the P1 virion. Cleavage of the protein at a position at or shortly after 421 would result in a ~46 kDa product, which matches closely the observed 50 kDa DdrB band. Presumably some small amount of the C-terminal fragment somehow remains with the virions, and this fragment would also be ~50 kDa and co- migrate with the C-terminal portion. The phenotype associated with ddrB deletion is an increase in protection against EcoB and EcoK restriction. One possibility is that the absence of DdrB allows for incorporation of more DarB into the virion, providing greater antirestriction activity. However, band densitometry does not convincingly show greater DarB band intensity in ddrB mutants (data not shown). An alternative hypothesis is that DdrB actively suppresses DarB-associated antirestriction activity. In

either case, the role of DdrB in the P1 antirestriction system is not clear, as its presence appears to counterintuitively hinder the fitness of P1 as measured by its ability to overcome host restriction.

As shown in Fig. 2.5, P1∆hdf, P1∆darA and P1∆ddrA virions all fail to incorporate

DarB, DdrB and Ulx. EcoB and EcoK restriction sensitivity of these mutants can be explained by the absence of DarB because DarB is most likely responsible for EcoB and

EcoK antirestriction activity as discussed above. However, hdf, darA and ddrA mutants are also all sensitive to EcoA restriction, which suggests that one of these three genes are responsible for the EcoA antirestriction activity of P1. Deletion of either hdf or darA results in virions missing both Hdf and DarA, which indicates that these proteins are co- dependent for virion incorporation. Both the P1∆hdf and P1∆darA virions are also missing DdrA in addition to DarB, DdrB and Ulx mentioned above. It was suggested that DarA is responsible for EcoA protection (Iida et al., 1987), however, DarA was found to be present in P1∆ddrA virions, which are sensitive to EcoA restriction (Figs.

2.5, 2.3C). This suggests that DarA is not solely responsible for protection from EcoA restriction as previously proposed. Aside from DarB, DdrB and Ulx, the only protein observed to be absent in P1∆ddrA is DdrA itself. Thus, DdrA may be the protein that protects P1 DNA from EcoA-mediated endonuclease activity, although it is not clear if

DarA and Hdf are also required for DdrA activity or are simply required for incorporation of DdrA into the virion.

Band densitometry was performed on SyproRuby-stained SDS-PAGE of P1 using

ImageJ (Schneider et al., 2012) to estimate the copy numbers of P1 antirestriction system components associated with the virion. These calculations suggest that there are

~45 DarB, ~100 DarA, ~40 DdrB and ~400 Hdf molecules in the average P1 virion

(Table 2.3). The copy number of Ulx and DdrA could not be estimated due to their low mass and band intensity. These values indicate that P1 capsids contain ~35 MDa of major capsid protein and a combined mass of ~28 MDa of Dar proteins (DarB, DarA,

DdrB and Hdf) per virion, indicating the Dar proteins contribute to a significant proportion of the mass of P1 heads.

Morphological defects are linked to hdf and darA

Under normal conditions, P1 is known to produce three morphological head size variants termed “big” (P1B, ~86 nm), “small” (P1S, ~65 nm) and “minute” (P1M, ~47 nm)

(Walker & Anderson, 1970). The P1B variant (hereafter referred to as “normal” head size or P1N) packages the complete genome and is infectious, while P1S and P1M are defective as they can package only ~40% and ~10% of the P1 genome, respectively

(Walker & Anderson, 1970). The proportion of P1S virions has been reported to vary depending upon the host, with P1 lysates induced from E. coli K-12 containing 20-37%

P1S heads; P1M heads have been reported as less than 1% of most lysates (Walker &

Anderson, 1970).

Table 2.3. Copy number of virion-associated proteins of P1 antirestriction system. Reprinted with permission from Piya et al., 2017. The copy number of constituent proteins of P1 antirestriction system was determined by densitometry on SyproRuby stained SDS-PAGE of P1 and mutants using ImageJ. The predicted copy number of tail sheath protein was determined based on the number of striations previously reported (six copies per striation, 54 striations) (Walker & Anderson, 1970) and used as an internal standard in all calculations. The molecular mass of tail sheath, DarB, and Hdf were determined from the predicted translational product of their respective genes. The molecular mass of DarA was determined based on the processing of product of darA into the mature form found in virions (Streiff et al., 1987). Finally, the molecular mass of major capsid protein (MCP) and DdrB were estimated based on their migration on SDS-PAGE. The copy number of MCP was also calculated from densitometry as a quality check; there are predicted to be 955 copies of the MCP per virion based on the previously reported T=16 P1 capsid symmetry (Walker & Anderson, 1970). The copy number of MCP as determined by densitometry is ~17% lower than expected, which provides some indication of the accuracy of this analysis. Data is shown for analyses of two replicate SDS-PAGE gels. Copy number per virion Molecular Total predicted protein Phage P1 proteins Mass (kDa) Repeat 1 Repeat 2 Average mass per virion (MDa)

Tail Sheath 57 324* 324* 324* 18 Major Capsid Protein 44 769 819 794 35 DarB 252 41 47 44 11 DarA 59 108 102 105 6 DdrB 50 40 34 37 2 Hdf 22 426 371 399 9 *: predicted copy number based on literature; this number was used as the basis for calculating copy number of the other proteins.

Fig. 2.7. Purification of virions using Cesium chloride isopycnic gradient centrifugation. Reprinted with permission from Piya et al., 2017. Parental P1 and mutants were purified by using isopycnic gradient of Cesium chloride as described earlier. The phage bands were extracted using a syringe and refractive index was measure using Abbe Refractometer. The refractive index was converted to density (g cc-1) by comparison with standard CsCl density-refractive index correlation table. The corresponding phage bands indicated with bold-face densities were used for comparative SDS-PAGE analysis shown in Fig. 5. The phage bands corresponding to grey-color densities denote tail-less phage heads. After dialysis, virions from all extracted bands were imaged under TEM and head size survey was conducted as described earlier. The major population of head-size variants present in each phage bands is indicated after density, separated by a colon (:). “N” denotes normal head size virions, “S” denotes small head size virions, and “M” denotes mixed population of normal and small head size virions.

In isopycnic gradient purification of P1 and its antirestriction mutants, variations in banding patterns were observed in some of the mutant phages, particularly P1Δhdf and

P1ΔdarA (Fig. 2.7). These mutants exhibited more intense bands at regions in the gradients corresponding to lighter buoyant densities than the parental P1, which suggested that the disruption of these genes also affected viral morphogenesis. The phage bands were extracted whenever possible and the head size of purified phages were measured from negative-stained TEM images and their distribution is described in Fig.

2.7.

In order to study the effects of antirestriction gene deletions on the overproduction of

P1S virions, the parental and mutant phages were induced from lysogens of E. coli

MG1655 and a head-size survey was conducted for each phage by measuring the diameters of several hundred phage heads photographed from negative-stain TEM images prepared from the phage lysates. Observed phage head diameters ranged from

52 nm to 95 nm (Fig. 2.8). In accordance with previous reports, head diameters >67 nm were classified as “normal” (i.e., P1N) and head diameters ≤67 nm were classified as small (P1S) variants. Virions with “minute” (P1M) head size were rarely observed and were excluded from classification. All phages produced heads of a bimodal size distribution, with a smaller size class of ~60 nm and a normal size class centered around

~80 nm (Fig. 2.8, Table 2.4). As shown in Fig. 2.8, the parental P1 produced virions with ~82% normal-sized heads and ~18% small heads, in agreement with previous observations of P1 (Walker & Anderson, 1970). In lysates of P1Δhdf and P1ΔdarA this relationship was essentially inverted, with P1Δhdf producing ~85 % small-headed virions and P1ΔdarA producing ~82% small-headed virions. Other phage mutants produced small heads at frequencies similar to the parental P1, ranging from 14.1% for

P1ΔddrB to 28.1% for P1ΔdarB, however this variation is within the range previously reported for P1 (Walker & Anderson, 1970).

Fig. 2.8. Head size variation in parental P1 and mutants. Reprinted with permission from Piya et al., 2017. Lysates of phage P1 contain virions of two major head size classes. The virions of >67 nm head diameter have been classified as “normal” and those of ≤ 67 nm as “small”. Panel A: Transmission electron micrograph (TEM) showing typical normal head (left) and small head (right) virions. Panel B: The diameters of parental P1 and antirestriction mutant virions were measured from TEM images. The X-axis in each histogram denotes the head size diameter in nm and the Y-axis denotes the percentage of virions of each head size present. The phage genotype (parental P1 or antirestriction mutant) and the number of virions measured are denoted in the upper right of each histogram. On the right, the percentages of virions falling into the small and normal head size classes for each phage mutant are summarized. The darB, ulx, ddrB and ddrA mutants have similar head size distributions as parental P1, while this distribution is reversed in hdf and darA mutants, which are dominated by small-headed virions.

Fig. 2.8 Continued.

Table 2.4. Mutations in phage P1 antirestriction genes affect head morphogenesis. Reprinted with permission from Piya et al., 2017. Lysates of phage P1 and its antirestriction mutants were observed by negative stain TEM and head diameters of the phage populations were measured. Phage heads were placed into two size classes, small (≤ 67 nm) and normal (> 67 nm). P1 hdf and darA mutants exhibited high proportions of small-headed virions. Phage No. virions Dimension of small head Dimension of normal heads strain measured (nm) (Average ± S.D.) (nm) (Average ± S.D.) P1 765 60 ± 3 79 ± 4 ∆darB 303 59 ± 3 78 ± 4 ∆ulx 301 60 ± 3 80 ± 4 ∆ddrB 290 61 ± 3 81 ± 3 ∆hdf 359 58 ± 3 77 ± 4 ∆darA 316 60 ± 3 78 ± 4 ∆ddrA 302 60 ± 4 80 ± 5

Previous work has shown that deletion of the entire darA operon, or transposon insertions within lydA and darA, resulted in overproduction of P1S virions in a phenotype denoted Vad- (for viral architecture determinant) (Iida et al., 1998). Based on the data presented here, the Vad- phenotype can be assigned to two genes within the darA operon, hdf and darA. These genes are directly adjacent to each other (Fig. 2.1B) and are the earliest non-lysis genes present in the Pdar transcript. Deletion of either gene results in the failure to incorporate any of the other virion-associated antirestriction components (Fig. 2.5), and the loss of antirestriction activity against EcoA, EcoB and

EcoK (Fig. 2.3). The presence of the Vad- phenotype in both P1Δhdf and P1ΔdarA indicates that both Hdf and DarA play important roles in determining head size during

P1 morphogenesis.

In many viruses (bacteriophages such as P22, T2,  and eukaryotic viruses such as HSV-

1 and other herpesviruses), assembly of the major capsid protein into phage heads of the correct size and shape is guided by a scaffolding protein, resulting in a spherical immature procapsid (Mateu, 2013). Studies in phages T4, P22 and  have shown that procapsids are assembled initially without any DNA (King et al., 1980). During procapsid maturation and concomitant DNA packaging, the scaffolding protein is lost from the capsid (Dokland, 1999). Absence of functional scaffolding proteins results in aberrant capsid assembly or no assembly at all (Dokland, 1999; Aksyuk & Rossmann,

2011). No major scaffold protein has been identified for P1 (Lobocka et al., 2004).

Disruption of hdf and darA results in an increased proportion of small P1 heads (Fig.

2.8), suggesting that Hdf and DarA play roles in directing proper capsid assembly, most likely at the time of procapsid formation. However, Hdf and DarA are still present in mature virions (Fig. 2.5) and hence do not fit the classical definition of scaffolding proteins which are present in viral proheads but absent in mature virions (Dokland,

1999). Moreover, the presence or absence of these proteins do not result in strict phenotypes: the deletion of hdf or darA still produces ~20% normal-sized phage heads, and the parental P1 containing intact hdf and darA produces ~20% aberrant capsids. Hdf and DarA may not play a direct role in the observed antirestriction activity of P1, but rather their absence prevents the incorporation of the other antirestriction system components.

Since the absence of either Hdf or DarA was found to induce greater production of small head-sized virions, we wished to determine if the small head-size progeny of parental P1 are capable of incorporating Hdf and DarA. Comparative SDS-PAGE band intensities of P1N and P1S virions suggest that small-head phages incorporate ~65% of the major capsid and ~40% of the DarB proteins relative to normal P1N virions (Fig. 2.9). These numbers are comparable to what would be expected from a reduction in head diameter from 80 nm to 60 nm, with concomitant reductions of capsid surface area by ~56% and capsid volume by ~42%. While the P1S virions are capable of incorporating both DarA and Hdf, band densitometry suggests that P1S heads incorporate only ~14% and ~12% of DarA and Hdf proteins, respectively, relative to P1N (Fig. 2.9). At this point it is difficult to determine if P1S heads are produced as a result of reduced incorporation of

Percent band intensities of Phage proteins in P1S relative to P1 P1N virions proteins Repeat Repeat Average 1 2 MCP 62 68 65 DarB 38 44 41 DarA 14 14 14 Hdf 16 8 12

Fig. 2.9. Comparative SDS-PAGE of P1N and P1S virions. Reprinted with permission from Piya et al., 2017. P1N, P1S and P1darA small head-size virions were extracted from isopycnic CsCl gradients, run on 12% Tris-glycine SDS-PAGE and stained with SyproRuby as described above. Band intensities in each lane were normalized to the tail sheath band and densitometry was performed on the major capsid protein (MCP), DarB, DarA and Hdf bands in ImageJ. As expected, bands corresponding to DarB, DarA, and Hdf were not observed in P1darA small head-size virions. This analysis was repeated twice and the relative band intensities measured in P1S virions are summarized in the table below the gel image.

Hdf and DarA, or if lower amounts of Hdf and DarA are incorporated into proheads that have already committed to the smaller size.

The incorporation of the P1 antirestriction proteins into the capsid appears to be linked to the early stages of capsid morphogenesis. All of the virions run on SDS-PAGE in Fig.

2.5 were taken from CsCl bands corresponding to normal phage buoyancy (1.46-1.48 g cc-1, Fig. 2.7) and electron microscopy surveys of these fractions confirmed that they contained >95% normal head size P1N virions (data not shown). Even though the

P1Δhdf and P1ΔdarA virion proteomes shown in Fig. 2.5 are the minority fraction of the phage lysate containing normal-sized heads, the downstream antirestriction components

DdrA, DdrB, DarB and Ulx are still absent from the virions, demonstrating that the formation of normal-sized heads is in itself not sufficient for incorporation of these antirestriction proteins. This suggests that it is Hdf and DarA themselves, and not their effects on head morphogenesis, that determine the incorporation of the other antirestriction proteins.

Conclusions

The P1 Dar system is composed of multiple proteins: Hdf, DarA, DdrA, DdrB, DarB and

Ulx. The role of Hdf and DarA in antirestriction system is unique because they are also essential for the production of normal head-size virions. In the presence of Hdf and

DarA, ~80% of P1 progeny are of normal head-size and ~20% are of small head-size

(Fig. 2.10A). The distribution of normal and small head-size is reversed in the absence

Fig. 2.10. Models of P1 head-size determination and antirestriction component incorporation. Reprinted with permission from Piya et al., 2017. Panel A: The parental P1 with intact darA and hdf produces ~80% normal head-size (P1N) and ~20% small head-size (P1S) progeny. In the absence of either Hdf or DarA, this ratio is inverted, producing progeny with mostly (~80%) aberrant small heads. Panel B: Incorporation of P1 antirestriction system components follows a distinct pathway. The pathway presumably begins with the initiation of prohead formation by the phage portal and scaffold proteins, followed by co-incorporation of Hdf and DarA into the prohead. In addition to exacerbating the head-size defect as shown in Panel A, the absence of Hdf or DarA prevents the incorporation of all downstream Dar components. The incorporation of Hdf and DarA does not confer protection against EcoA, EcoB or EcoK restriction. Incorporation of DdrA appears to be the next step, as ddrA mutants incorporate only DarA and Hdf. Incorporation of DdrA provides protection against EcoA restriction. The incorporation of DarB, DdrB and Ulx is difficult to separate into separate steps, but packaging of DarB into virions provides protection against EcoB and EcoK restriction. Ulx appears to be an enhancer of DarB packaging as ulx mutants incorporate less DarB than normal, while DdrB acts as a negative regulator of DarB activity.

Fig. 2.10 Continued.

of either Hdf or DarA (Fig. 2.8). Because the absence of either Hdf or DarA results in a high abundance of aberrant small capsids, it is probable that these proteins play a role in the early stages of prohead formation. The proteins Hdf, DarA and DdrA are required for the protection of phage DNA from restriction by the EcoA Type I R-M system, and

DarB is required for protection against EcoB and EcoK. Ulx enhances the antirestriction phenotype of DarB, and DdrB appears to negatively affect this activity by an unknown

mechanism (Fig. 2.10B). All of these phenotypes could be complemented in trans.

Proteomic analysis of isogenic P1 mutants shows a clear order of incorporation for each component into the mature virion, as illustrated in (Fig. 2.10B). In this proposed model,

Hdf and DarA are incorporated first in a co-dependent manner, shortly after prohead initiation. The absence of either Hdf or DarA results in failure to incorporate any of the other antirestriction proteins. Following Hdf and DarA, DdrA is incorporated, followed by DarB, DdrB and Ulx. The incorporation of these last three proteins are not strictly co-dependent: the absence of DarB results in no apparent Ulx incorporation, but absence of Ulx results only in reduced DarB incorporation. This behavior suggests that Ulx acts as a chaperone or packaging factor to increase the amount of DarB incorporated into the virion. The presence of DdrB results in reduced antirestriction activity, presumably by negatively affecting DarB but the mechanism is not known.

Given the incorporation of these proteins into the P1 virion and the cis-acting nature of the antirestriction phenotype, it is evident that the P1 antirestriction system must exert its activity upon infection of the host cell (Iida et al., 1987). Since it is difficult to imagine how these proteins could act while still in the confines of the capsid, they must be introduced into the host cytoplasm via the phage tail upon infection. It is not a requirement that all six of the antirestriction proteins described here be translocated into the cytoplasm to perform some direct protective action. Some proteins, particularly the more upstream components such as DarA and Hdf, likely only serve to ensure the incorporation of the active components into the virion. Strictly speaking, a minimum of

only two proteins are required to account for the observed phenotypes: one that provides protection against EcoA, and one that protects against EcoB and EcoK. Based on bioinformatic evidence, DarB is the likely candidate to provide EcoB and EcoK protection as the predicted methyltransferase and helicase domains of DarB suggest a possible enzymatic mechanism to provide protection from host restriction. The protein or proteins responsible for the observed anti-EcoA activity are more difficult to assign.

DdrA is one candidate, as phage lacking DdrA contain DarA and Hdf but do not express any antirestriction activity.

The translocation of phage proteins from the head into the host cytoplasm is a known feature in several Caudovirales phages. The capsid of the temperate Salmonella

Typhimurium podophage P22 contains three internal proteins, gp16, gp20, and gp7, that leave the confines of the capsid to perform their function of assisting in translocation of

DNA across host membranes (Jin et al., 2015). Coliphage N4 is known to translocate its

~380 kDa DNA-dependent RNA polymerase from the phage head into the host cytoplasm early in the infection process (Choi et al., 2008). Perhaps the closest analog to the P1 system shown in this work is the paradigm coliphage T4, whose heads contain three non-essential internal proteins, IPI, IPII and IPIII (Black & Ahmad-Zadeh, 1971).

The ~10 kDa T4 IPI is ejected into the host cytoplasm during phage infection and protects the incoming phage DNA by inhibiting a glucosyl-hmC DNA-specific restriction endonuclease (Bair et al., 2007). The proteins of the Dar system are assembled into the P1 capsid and at least some of them are ejected into the host

cytoplasm, where they protect P1 DNA from cleavage by Type I R-M systems. In our study of the P1 antirestriction system, we determined that the incorporation of the P1 antirestriction proteins follow a distinct pathway that involves at least six proteins. We also found that proteins involved in the assembly pathway influence head morphogenesis. Phage P1 is the first reported instance, to our knowledge, where the antirestriction system has also been implicated in capsid morphogenesis.

CHAPTER III

CHARACTERIZATION OF P1 GENES OF UNKNOWN FUNCTION AND

PACKAGING OF DARB INTO P1 PROCAPSIDS

Introduction

Bacteriophages (phages) are natural predators of bacteria. The constant evolutionary arms race between bacteria and phages has led to development of diverse defense and counter-defense strategies against each other (Labrie et al., 2010). Caudovirales phages such as P1 initiate the infection cycle by adsorption to host receptors, followed by ejection of DNA into host cytoplasm. Bacterial restriction and modification (R-M) systems provide a barrier to foreign DNA, including phage DNA, by cleaving inappropriately modified DNA. Based upon subunit stoichiometry, DNA specificity and restriction mechanisms, R-M systems are broadly categorized as type I, II and III.

(Kruger & Bickle, 1983; Loenen et al., 2014). Please refer to Chapter I of this dissertation for detailed information on mechanisms of action of restriction-modification systems.

To avert DNA cleavage by the bacterial R-M systems, phages have evolved with diverse antirestriction mechanisms (Tock & Dryden, 2005). Enterobacteriaceae phage P1 protects its DNA from type I R-M system mediated DNA cleavage by injecting Dar proteins into host. Earlier reports indicated that Dar system was composed of two proteins, DarA and DarB, which are encoded in two distinct operons in P1 genome (Iida

et al., 1987; Lobocka et al., 2004). It was determined from plating assays that P1 darA mutants are severely restricted in host with type I EcoA, whereas P1 darB mutants are severely restricted in host with type I EcoB or EcoK systems (Iida et al., 1987). Recent study has shown that several gene in darA and darB operons, including darA and darB themselves, encode components of P1 antirestriction system. P1 mutants deleted for ulx show darB-like restriction phenotype, whereas mutants deleted for hdf or ddrA show darA-like phenotype (Fig. 3.1). Moreover, Hdf and DarA, which were shown to be required for incorporation of all Dar proteins, were also found to affect capsid size determination (Piya et al., 2017). Since multiple genes in two distinct operons comprise the Dar system, we hypothesized that P1 genes from other different operons might influence P1 antirestriction system. We also interested to see if any other P1 genes could affect capsid size determination.

Genetic evidence has established that DarB is required for protection of P1 DNA from type I EcoB and EcoK systems following ejection of DNA into host (Iida et al., 1987;

Piya et al., 2017). Presence of DarB in mobile genetic elements suggest that DarB could protect these elements from restriction during their transfer across different hosts in nature (Gill et al., 2011). Biological function of DarB has been studied in context of phage P1 only. However, little is known about the assembly of DarB into P1 procapsids and biochemical activity of DarB leading to protection of DNA.

P1 Dar system is encoded in two operons, separated by genes of unrelated functions, thus it is possible that other as-yet undetected genes may also play a role in this

Fig. 3.1. Genome map of P1. Rectangular blocks in the figure represent P1 genes. The genes above the black line are transcribed towards right, whereas those below the black line are transcribed towards left. The genes darB, ulx, hdf, darA, ddrA, and ddrB encoding known components of P1 antirestriction are indicated. P1 genes of unknown function that were knocked out in this study are colored red or grey. P1 mutants knocked out for genes in red color did not form plaques and were not tested for plating efficiency. P1 mutants knocked out for genes in grey color formed plaques and were lysogenized into E. coli strain WA921. Thermally induced lysates were tested for plating efficiency in E. coli strains with type I EcoA, EcoB or EcoK system. The genes encoding major structural components are indicated for reference. The ruler denotes the nucleotide position of genes as in the published P1 genome (NC_005856.1).

antirestriction system. In order to identify other components of the P1 antirestriction system, if any, a library of P1 mutants knocked out for genes of unknown function was constructed. In order to further examine the incorporation of Dar components into mature virions, P1 mutants defective for DNA packaging was constructed. Several truncations of DarB were also generated to study mechanisms of its incorporation into

P1 capsids. Moreover, we have optimized purification of DarB to elucidate reaction mechanisms in vitro.

Materials and methods

Bacterial strains and phages

The bacterial strains and phages used in this study are listed in Table 3.1. All phage P1 mutants used in this study are isogenic gene deletions of P1CMclr100, simply referred to as P1 (Rosner, 1972). Unless specified otherwise, E. coli strains, were cultured in LB broth or LB agar at 37 °C as described before (Piya et al., 2017). Culture was grown at

30 °C to maintain P1 lysogen and shifted to 42 °C to thermally induce P1. As needed, the medium was supplemented with 10 µg mL-1 chloramphenicol (LB cm), 30 µg mL-1 kanamycin (LB kan) or 100 µg mL-1 ampicillin (LB amp).

Generation of single gene P1 knockout mutants

Single-gene deletions knockout library of P1 was constructed by using  Red recombinase mediated homologous recombination method as described before

(Datsenko & Wanner, 2000). In brief, P1 was lysogenized into E. coli

Table 3.1. Bacterial strains, phages and plasmids

Strains, phages or Genotype or relevant characteristic Reference/source plasmids E. coli strains - - - - - WA2379 leu met lac rA mA (Arber & Wauters- Willems, 1970) / The Coli Genetic Stock Center, Yale University (CGSC) - - + + W3110 F  rpoS(Am) rph-1 Inv(rrnD-rrnE) rK mK (Iida et al., 1987; Hayashi et al., 2006) / CGSC ------WA921 thr leu met lac rK mK (Wood, 1966; Arber & Wauters- Willems, 1970) / CGSC + - - - + + WA960 thrB leu met lac rB mB (Wood, 1966; Arber & Wauters- Willems, 1970) / CGSC BW25113(pKD46) F- l- rpoS(Am) rph-1 rrnB3 ΔlacZ4787 hsdR514 Δ(araBAD)567 (Baba et al., 2006) Δ(rhaBAD)568 / CGSC q BW25141(pKD4) lacI rrnBT14 ΔlacZWJ16 ΔphoBR580 hsdR514 ΔaraBADAH33 (Datsenko & + ΔrhaBADLD78 galU95 endABT333 uidA(ΔMluI)::pir recA1 Wanner, 2000) / CGSC

MG1655 lacIq1tonA::Tn10 (Park et al., 2006) / Lab stock

Phages

P1ΔdarA in-frame deletion of darA in P1CMclr100 (Piya et al., 2017) P1ΔdarB in-frame deletion of darB in P1CMclr100 (Piya et al., 2017) P1ΔdarAΔpacA in-frame deletion of pacA in PΔdarA This study P1ΔdarBΔpacA in-frame deletion of pacA in PΔdarB This study P1Δprt in-frame deletion of prt in P1CMclr100 This study P1Δpro in-frame deletion of pro in P1CMclr100 This study P1ΔpmgA in-frame deletion of pmgA in P1CMclr100 This study P1ΔpmgB in-frame deletion of pmgB in P1CMclr100 This study P1ΔpmgC in-frame deletion of pmgC in P1CMclr100 This study P1ΔpmgF in-frame deletion of pmgF in P1CMclr100 This study P1ΔpmgG in-frame deletion of pmgG in P1CMclr100 This study P1ΔupfA in-frame deletion of upfA in P1CMclr100 This study P1ΔupfB in-frame deletion of upfB in P1CMclr100 This study P1ΔupfC in-frame deletion of upfC in P1CMclr100 This study

Table 3.1. Continued. Strains, phages or Genotype or relevant characteristic Reference/source plasmids P1Δuhr in-frame deletion of uhr in P1CMclr100 This study P1Δupl in-frame deletion of upl in P1CMclr100 This study P1Δdbn in-frame deletion of dbn in P1CMclr100 This study P1ΔpmgL in-frame deletion of pmgL in P1CMclr100 This study P1ΔpmgM in-frame deletion of pmgM in P1CMclr100 This study P1ΔpmgN in-frame deletion of pmgN in P1CMclr100 This study P1ΔpmgO in-frame deletion of pmgO in P1CMclr100 This study P1ΔpmgP in-frame deletion of pmgP in P1CMclr100 This study P1Δppp in-frame deletion of ppp in P1CMclr100 This study P1ΔpmgQ in-frame deletion of pmgQ in P1CMclr100 This study P1ΔpmgR in-frame deletion of pmgR in P1CMclr100 This study P1ΔpmgS in-frame deletion of pmgS in P1CMclr100 This study P1Δpap in-frame deletion of pap in P1CMclr100 This study P1ΔpmgT in-frame deletion of pmgT in P1CMclr100 This study P1ΔpmgU in-frame deletion of pmgU in P1CMclr100 This study P1ΔpmgV in-frame deletion of pmgV in P1CMclr100 This study P1ΔupfM in-frame deletion of upfM in P1CMclr100 This study P1ΔupfN in-frame deletion of upfN in P1CMclr100 This study P1ΔupfO in-frame deletion of upfO in P1CMclr100 This study P1ΔpdcA in-frame deletion of pdcA in P1CMclr100 This study P1ΔpdcB in-frame deletion of pdcB in P1CMclr100 This study

Plasmids (Cherepanov & + ts R R pCP20 FLP+,  cI857 ,  pR Rep , Ap , Cm Wackernagel, 1995) / Zeng Lab pdarB P1 darB cloned into pBAD24 (Piya et al., 2017) P1 darB fragment cloned into pBAD24 to express truncated DarB pdarB 1-1750 This study residues 1-1750 P1 darB fragment cloned into pBAD24 to express truncated DarB pdarB 90-1750 This study residues 90-1750 P1 darB fragment cloned into pBAD24 to express truncated DarB pdarB 90-2255 This study residues 90-2255

Table 3.1. Continued. Strains, phages or Genotype or relevant characteristic Reference/source plasmids P1 darB fragment cloned into pBAD24 to express truncated DarB pdarB 5-2255 This study residues 5-2255 P1 darB fragment cloned into pBAD24 to express truncated DarB pdarB 10-2255 This study residues 10-2255 P1 darB fragment cloned into pBAD24 to express truncated DarB pdarB 20-2255 This study residues 15-2255 P1 darB fragment cloned into pBAD24 to express truncated DarB pdarB 25-2255 This study residues 20-2255 P1 darB fragment cloned into pBAD24 to express truncated DarB pdarB 25-2255 This study residues 25-2255 Addgene / Lab pMB838 source of gentamicin resistance cassette stock pBAD24g pBAD24 with gentamycin resistance cassette This study ppro P1 pro cloned into pBAD24g This study ppmgA P1 pmgA cloned into pBAD24g This study ppmgB P1 pmgB cloned into pBAD24g This study ppmgC P1 pmgC cloned into pBAD24g This study ppmgG P1 pmgG cloned into pBAD24g This study ppmgR P1 pmgR cloned into pBAD24g This study Millipore Sigma / pETDuet-1 Protein expression vector Lab stock pETDuet_darB P1 darB cloned into pETDuet-1 This study pETDuet_2x- pETDuet_darB engineered to introduce additional N-terminal His- This study His_darB tag pBAD24_2x- gene fragment encoding 2x-His_DarB cloned into pBAD24 This study His_darB pFTSKi-tetR- (Shao et al., 2017) template for PCR amplification of tetR-mCherry mCherry / Zeng Lab Derivative of pBAD24 constructed to express recombinant proteins pBAD24_darBN9 with N-terminal 9 residues to DarB fused to N-terminus of any This study protein Derivative of pBAD24 constructed to express recombinant proteins pBAD24_darBN3 with N-terminal 30 residues to DarB fused to N-terminus of any This study 0 protein pBAD24_darBN9 tetR-mCherry cloned into pBAD24_darBN9 This study _tetR-mCherry pBAD24_darBN3 tetR-mCherry cloned into pBAD24_darBN30 This study 0_tetR-mCherry

BW25113(pKD46). P1 lysogens were grown in LB cm + amp. At OD550 = 0.1, 1 mM L- arabinose was added to induce Red proteins and cells were cultured up to OD550 0.5-0.6.

At this point, the culture was harvested, and cells were made electrocompetent. Cells were transformed with gel purified DNA (QIAquick Gel Extraction Kit), which was obtained by PCR amplification of EcoP1R FRT-flanked kan gene from plasmid pKD4.

The primers to amplify kan gene were designed to include homology regions to P1 DNA and pKD4 priming sites, as described before. The transformed cells recovered in LB at

30 °C for 2 h. After overnight incubation at 30 °C, CmR and kanR colonies were selected, and mutations were confirmed by PCR and sequencing (Piya et al., 2017). The primers used to amplify kan inserts are listed in Table 3.2.

Flp-mediated excision of kan insert and generation of double-gene knockout mutants

All single-gene deletion mutants of P1 were generated by replacing respective genes with kan gene flanked with FRT (Flp recombination target) sites as described above.

Saccharomyces cerevisiae encoded Flp enzyme mediates recombination between these

FRT sites, enabling excision of the antibiotic resistance gene (Cherepanov &

Wackernagel, 1995). E. coli MG1655 strain lysogenized with either P1ΔdarA or

P1ΔdarB was made electrocompetent and transformed with plasmid pCP20 as previously described (Cherepanov & Wackernagel, 1995; Datsenko & Wanner, 2000).

Lysogens of either P1ΔdarA or P1ΔdarB mutants are CmR and KanR, and plasmid pCP20 transformants are CmR and AmpR. P1ΔdarA or P1ΔdarB lysogens transformed with pCP20 were selected as CmR and AmpR colonies. The FLP gene can be induced

Table 3.2. Primers and synthetic DNA fragments Primers Forward primer 5'-3' Reverse primer 5'-3' Usage atcggacgtcatgttacgcagcag atcggagctcttaggtggcggt To PCR amplify gentamicin caacgatg acttgggtc resistance cassette from pMB838 atcggagctcctgtcagaccaagt atcggacgtcactcttcctttt To PCR amplify pBAD24 backbone ttactcatatatactttagattg tcaatattattgaag atactgagctcaatgaacaagcta atactggcgcgccgtcacggtc To clone darB into pETDuet-1 tctatgggggtgt aacgagacgac tgagcggataacaattcccctcta aatttcgattatgcggccgtgt To PCR amplify synthetic DNA g ac fragments designed to introduce additional His-tag to pETDuet-1 atcgtctagagtaaaggaggtatc atcgaagcttgtcacggtcaac To clone darB fused with 2x His-tag gatatgggcagcagccatcac gagacgac from pETDuet_darB-12His into pBAD24 atcgtctagagtaataaaggaggt atcgaagcttttatgagcgatc To clone darB into pBAD24 to atcgagatgaacaagctatctatg gccgataacaag express residues 1 - 1750 ggggtg atcgtctagagtaataaaggaggt atcgaagcttttatgagcgatc To clone darB into pBAD24 to atcgagatggaatattacacacca gccgataacaag express residues 90 - 1750 aagccgatcg atcgtctagagtaataaaggaggt atcgaagcttttatgcgtattg To clone darB into pBAD24 to atcgagatggaatattacacacca ttggatgacggc express residues 90 - 2255 aagccgatcg atcgtctagagtaataaaggaggt atcgaagcttttatgcgtattg To clone darB into pBAD24 to atcgagatgtctatgggggtgttt ttggatgacggc express residues 5 - 2255 cgctg atcgtctagagtaataaaggaggt atcgaagcttttatgcgtattg To clone darB into pBAD24 to atcgagatgcgctgttcaagtgtc ttggatgacggc express residues 10 - 2255 agcg atcgtctagagtaataaaggaggt atcgaagcttttatgcgtattg To clone darB into pBAD24 to atcgagatgagcgaaatattgaaa ttggatgacggc express residues 15 - 2255 tacattagggc atcgtctagagtaataaaggaggt atcgaagcttttatgcgtattg To clone darB into pBAD24 to atcgagatgtacattagggcaata ttggatgacggc express residues 20 - 2255 acatctcaccg atcgtctagagtaataaaggaggt atcgaagcttttatgcgtattg To clone darB into pBAD24 to atcgagatgacatctcaccgagcg ttggatgacggc express residues 25 - 2255 cc tttcgttaccctcgcgtcaaaaag acgtgggtccgttacgcaactt To knockout pacA from P1 genome agtttttacgaaaggaagcataag ctggccattaatcatcatcccc tggtgtaggctggagctgcttc gtacagatgggaattagccatg gtcc tccgcacgtatctgattgattg gtcttcccaaacaacacatcag Sequencing primers to verify pacA c deletion in P1 genome

Table 3.2. Continued.

Forward primer 5'-3' Reverse primer 5'-3' Usage aaattccgttcaaacacgatgtga tcagtgaccgttttcaaaacat To knockout prt from P1 genome attattctaattaaggtgcaatct cagtcattatcgtttccctctt tggtgtaggctggagctgcttc taaagaatgggaattagccatg gtcc ccgtatttaatcggcgctcg ggcgaccgtgaccgtaatag Sequencing primers to verify prt deletion in P1 genome aaatccttgaatcggtctttaaag tttggtggttatttaaacggat To knockout pro from P1 genome agggaaacgataatgactgatgtt tgattgaattattaaacgtgat ttgtgtaggctggagctgcttc gatgctatgggaattagccatg gtcc ttacctgttcagcgatgtgc gctattgtatgcagggcttc Sequencing primers to verify pro deletion in P1 genome tcaattgatgatccgttgatgaat ctgtctttagtcggtgttgttt To knockout pmgA from P1 gactacgcgagagttgattgatgg tactcatagcaccacgtcctgt genome ccgtgtaggctggagctgcttc gtgataatgggaattagccatg gtcc tgaaacgaagcggctgtttg gatataaagccttcagccagc Sequencing primers to verify pmgA deletion in P1 genome gcttcccttctgggctttagtttt ggaaaaacgcataggattcagt To knockout pmgB from P1 tctctttgaattaaggagcaagga gagcctttcagcgtcctgaatc genome tggtgtaggctggagctgcttc ggtataatgggaattagccatg gtcc cttccggcttaaccaaactg tcccgaatacctgcaagctc Sequencing primers to verify pmgB deletion in P1 genome gtagaatcggttaacacaccagat aagcaacattaagccccatatc To knockout pmgC from P1 tctacgaggtttcaatgacaccac agccccctcacttcaacatgga genome gagtgtaggctggagctgcttc tgagaaatgggaattagccatg gtcc aacacaacgcccattaaagg tcaacttcgacagagaactgc Sequencing primers to verify pmgC deletion in P1 genome ttttttgtgcttaaatcatgtcaa gataattccgggagagcggttg To knockout pmgF from P1 tatagcgaaattttgagcatatta ggttttttcatgcttttctgcg genome tggtgtaggctggagctgcttc cagagaatgggaattagccatg gtcc ttagtaccgaagatgtgactgctg agctgaacccgacacatattgc Sequencing primers to verify pmgF deletion in P1 genome ggaactccggcctttaacttgaat agtgttattgtgccccatataa To knockout pmgG from P1 ggctcctatagcttatggggttta aatccttttactggaacgcccg genome cagtgtaggctggagctgcttc aacaatatgggaattagccatg gtcc aggttgttcgtaacggtgag ttcaggaatttgcgacgtgc Sequencing primers to verify pmgG deletion in P1 genome

Table 3.2. Continued

Forward primer 5'-3' Reverse primer 5'-3' Usage gttaatgattacaaccgagctatt ccaacgtagccggttggaaggg To knockout upfA from P1 genome agcggtaactaaaagggattttta agtcgtattattctacgctttc tggtgtaggctggagctgcttc gatgagatgggaattagccatg gtcc aatttaacgccgacgccgac tttccctccagcacacatcg Sequencing primers to verify upfA deletion in P1 genome gagacggcctagttcaggtaagtt tgaagaaaaattatcaatgaag To knockout upfB from P1 genome aggagatatcatgctggaaaaaga tcctttgttactgtgccgcttt ctgtgtaggctggagctgcttc gtttaaatgggaattagccatg gtcc tcccgtagagagcgaaccac ggcttgctttagctgatgacg Sequencing primers to verify ufpB deletion in P1 genome aatctaagttaaaataacgaaaat cgtctccggctgcaacggtaga To knockout upfC from P1 genome cagagcaaatcattggtgatgacg gccgcatgaaacaggatcacca tggtgtaggctggagctgcttc acgcatatgggaattagccatg gtcc tgcatctgacgactccaaac gctggtaatagtgtcgggatag Sequencing primers to verify upfC deletion in P1 genome accattcagccatcgcccttcaat atagtaatgtttttccagtgct To knockout uhr from P1 genome gggcatttgtttggagtcgtcaga taaacatgttgtaacccttgaa tggtgtaggctggagctgcttc tatcaaatgggaattagccatg gtcc atacttgccacgtcatcacc tttgtttcgcgctttgcttc Sequencing primers to verify uhr deletion in P1 genome ttcaccgcaacggaaagagcattc tggtgaatgcacaggctgatgt To knockout upl from P1 genome ctggtggacctgtagattgggata gccgcaactacagtagtgcgcg tggtgtaggctggagctgcttc ctttgcatgggaattagccatg gtcc gggaaataaggttgcggtgc cgcaggaacgaccaataaacg Sequencing primers to verify upl deletion in P1 genome cggtttgataactatgccggtgct ggcttttctgttatgacgggtt To knockout dbn from P1 genome gactggcaagaggattattaatgc caattttttatccgttaccgcg aagtgtaggctggagctgcttc cgacggatgggaattagccatg gtcc aggacgcagacctgattatg gcattcatctcttccgcagg Sequencing primers to verify dbn deletion in P1 genome aaatgtatcattctgcccttaagt aaactcttcctcagtaaagcgc To knockout pmgL from P1 aggttcttcacgaggaaacaaaat aacataatcagtcaaatcctgc genome tggtgtaggctggagctgcttc cggtcgatgggaattagccatg gtcc gtgctttgggttgtagtgattg agtgctcgcaatttccgtcg Sequencing primers to verify pmgL deletion in P1 genome

Table 3.2. Continued.

Forward primer 5'-3' Reverse primer 5'-3' Usage gcaaaacagcgtaaatatggcatg agcgcacatgcactaattaatt To knockout pmgM from P1 cgaccggcaggatttgactgatta tattattttaagcagcatacaa genome tggtgtaggctggagctgcttc ccacttatgggaattagccatg gtcc cagaaagcaggcacaggtcg aggcgagcattcatttcagc Sequencing primers to verify pmgM deletion in P1 genome catgtgcgctctttgtggtagtgc agctgcaaattcagcctccagg To knockout pmgN from P1 actttaacatcgggagaataatcg cgagcattcatttcagcgatta genome tggtgtaggctggagctgcttc cagggtatgggaattagccatg gtcc gtgcagaacaagcgattacc ggaatctccctcccggtatg Sequencing primers to verify pmgN deletion in P1 genome gctgaatttgcagctgagaatgaa ctgcctctaaatttattcttgg To knockout pmgO from P1 cataccacccagggcgactaggac gaaacattgaattgcactgctc genome tggtgtaggctggagctgcttc ctacttatgggaattagccatg gtcc cgataagtggttgtatgctgc gaagcacagaacgccatctc Sequencing primers to verify pmgO deletion in P1 genome gcggagtgccgattatagcggtca gtgggtgctatcatgtattctt To knockout pmgP from P1 atggggccatcgactggcttaacc cagccattctttaagagtcatc genome gcgtgtaggctggagctgcttc tgcggaatgggaattagccatg gtcc cggctatgactgttctctgc gcaatagttccagacattcgac Sequencing primers to verify pmgP deletion in P1 genome ggtattccgcagatgactcttaaa ccagagcgaagaactaaagcaa To knockout ppp from P1 genome gaatggctgaagaatacatgatag tcttcatgctgcaccatcacct cagtgtaggctggagctgcttc ttcactatgggaattagccatg gtcc tgaacctggctttccatatcg tatccggtcggaacaactcg Sequencing primers to verify ppp deletion in P1 genome ggaaacctgacattgatgaaagtg tttaaaaatatggaattattag To knockout pmgQ from P1 aaaggtgatggtgcagcatgaaga agcaatattattctgattctcg genome ttgtgtaggctggagctgcttc ctcaaaatgggaattagccatg gtcc caccgggcaagaaactatcg gcgatacctggtgatgacatg Sequencing primers to verify pmgQ deletion in P1 genome atgtacactcatcacgttttttat ttgattggttttgtgctgaagt To knockout pmgR from P1 tagagcaatctacaaggtgcacta gatcatttcaaagttccgtatt genome tggtgtaggctggagctgcttc agcttgatgggaattagccatg gtcc ggccgatgttgtggaacctg gcatctccgctgactgaacc Sequencing primers to verify pmgR deletion in P1 genome

Table 3.2. Continued. Forward primer 5'-3' Reverse primer 5'-3' Usage atacttaggaaaaatgaccgaagc caccagcattaaaagtgacact To knockout pmgS from P1 acaagctaatacggaactttgaaa gtaactatcagcgaacgtaaat genome tggtgtaggctggagctgcttc agtgccatgggaattagccatg gtcc ggctgatcgcctttgttacg atgcagcctgtcttccgttg Sequencing primers to verify pmgS deletion in P1 genome ggtggagtgcgcccaccagcattt gttttaaaaatcaagatttatt To knockout pap from P1 genome ttttcgtccaatgaggagggcatt agagcaattattgttgatgaag tggtgtaggctggagctgcttc aagcgcatgggaattagccatg gtcc cgaattgggtttagaggaaacagc tttcacttcgcctttcatgc Sequencing primers to verify pap deletion in P1 genome tctctatcacttttggcggcatcg aatgtccggatagttgccagcc To knockout pmgT from P1 tcgcgcatgggtaagggggatgca tcgtaacccagggattagcact genome tggtgtaggctggagctgcttc ccagctatgggaattagccatg gtcc acgttaatggtcagggagtatg cgtgcgtctcatctgatctc Sequencing primers to verify pmgT deletion in P1 genome ttattacgccagatcatcataaac aggtgtttgatgtgatctgcaa To knockout pmgU from P1 aagccgagaaaagtgacctagaaa ctctcatacttcaccctcgctt genome gggtgtaggctggagctgcttc gtatcgatgggaattagccatg gtcc ggcatcacgcattctgttgg cccggttacgtctgttcttg Sequencing primers to verify pmgU deletion in P1 genome ttgccgttggtggatgctggcgat ttctgtatgtgctggtgggtac To knockout pmgV from P1 acaagcgagggtgaagtatgagag ctgtagttcagctttcgttggc genome ttgtgtaggctggagctgcttc atttaaatgggaattagccatg gtcc cacccagcaattacgagaaag cgcctgttattagccagattc Sequencing primers to verify pmgV deletion in P1 genome gatgaggataatagccagaatctg tgccgtaagctcacgttaacga To knockout upfM from P1 genome gctaataacaggcgcatctaaaaa ctttctttcaccgaatccaact tggtgtaggctggagctgcttc atataaatgggaattagccatg gtcc gtacccaccagcacatacag ttatctgccttgccgtgc Sequencing primers to verify upfM deletion in P1 genome aaacccaacccttatatagttgga acaaagttatgcacttgcaaga To knockout upfN from P1 genome ttcggtgaaagaaagtcgttaacg gggccattttctaaatattgtg tggtgtaggctggagctgcttc atgtttatgggaattagccatg gtcc tggctaataacaggcgcatc ccttatcccgttcttcctgac Sequencing primers to verify upfN deletion in P1 genome

Table 3.2. Continued.

Forward primer 5'-3' Reverse primer 5'-3' Usage gaaacatcacaatatttagaaaat tcccatccagaataattgagta To knockout upfO from P1 genome ggccctcttgcaagtgcataactt acgactattatttaaccagcaa tggtgtaggctggagctgcttc agtaacatgggaattagccatg gtcc tggattcggtgaaagaaagtcg gcagattctctggttgctcg Sequencing primers to verify upfO deletion in P1 genome tctgtcgccgacacgttacgtaga acatcacacctttaatcactga To knockout pdcA from P1 genome ttgtatggttctgcggagtagatt ttgggctttatctgctgcccgg aagtgtaggctggagctgcttc cattctatgggaattagccatg gtcc aagtctccgccacctacctg ccgtcttccatgtgctcgatta Sequencing primers to verify pdcA ac deletion in P1 genome gaatgccgggcagcagataaagcc taaagaaatagcaatacattag To knockout pdcB from P1 genome caatcagtgattaaaggtgtgatg agcaattttatctaacactcga tggtgtaggctggagctgcttc cgaatgatgggaattagccatg gtcc gtcccgtgaagcgttaaagatg agaagagatttagtgcggatca Sequencing primers to verify pdcB tg deletion in P1 genome tctagagtaaaggaggatcgatct atcgaaagctttcagtcattat To clone prt into pBAD24g tggcagacaataaaatcacgctat cgtttccctctttaaag c atcgatctagagtaaaggaggatc atcgaaagcttttatttaaacg To clone pro into pBAD24g gatcatgactgatgttttgaaaac gattgattgaattattaaacgt ggtcac gatg ggggatcctctagaataaaggagg aaaacagccaagctttcatagc To clone pmgA into pBAD24g tcagtacatggccaataataacga accacgtcctg aattgatcc ggggatcctctagaataaaggagg aaacagccaagctttcagcgtc To clone pmgB into pBAD24g gatgctaatgcttttacccctttt ctgaatcgg ccc ggggatcctctagaataaaggagg aaacagccaagctttcacttca To clone pmgC into pBAD24g gatgctaatgacaccacgacaatt acatggatgagaaaag actc

ggggatcctctagaataaaggagg aaacagccaagcttttactgga To clone pmgG into pBAD24g gatgctaatggctcctatagctta acgcccgaa tggg ggggatcctctagaataaaggagg aaacagccaagctttcaaagtt To clone pmgR into pBAD24g gatgctaatgtggccattccgacg ccgtattagcttgtg

Table 3.2. Continued

Forward primer 5'-3' Reverse primer 5'-3' Usage actctctactgtttctccataccc tgtatcaggctgaaaatcttct To PCR amplify synthetic DNA g ctcatc fragments designed to introduce N- terminal DarB residues into pBAD24 atcgagaattcgtgtctagattag atcgaggtaccttacttgtaca To clone tetR-mCherry into ataaaagtaaagtgattaacagcg gctcgtccatgc pBAD24_darBN9 and pBAD24_darBN30

Synthetic DNA fragments (5' - 3') aattgtgagcggataacaattcccctctagaaataattttgtttaact Synthetic DNA fragment to ttaagaaggagatataccatgggcagcagccatcaccatcatcaccac introduce additional His-tag to ggcagcagccatcaccatcatcaccacagccaggatccgaattcgagc pETDuet-1 tcggcgcgcctgcaggtcgacaagcttgcggccgcataatgcttaagt cgaacagaaagtaatcgtattgtacacggccgcataatcgaaattaat acgactcactata gacgctttttatcgcaactctctactgtttctccatacccgttttttt Synthetic DNA fragment to gggctagcaggaggtaatacaccatgaacaagctatctatgggggtgt introduce 9 N-terminal DarB ttgaattcaccatggtacccggggatcctctagagtcgacctgcaggc residues into pBAD24 atgcaagcttggctgttttggcggatgagagaagattttcagcctgat acagattaaatc gacgctttttatcgcaactctctactgtttctccatacccgttttttt Synthetic DNA fragment to gggctagcaggaggtaatacaccatgaacaagctatctatgggggtgt introduce 30 N-terminal DarB ttcgctgttcaagtgtcagcgaaatattgaaatacattagggcaataa residues into pBAD24 catctcaccgagcgccggaattcaccatggtacccggggatcctctag agtcgacctgcaggcatgcaagcttggctgttttggcggatgagagaa gattttcagcctgatacagattaaatc

from pCP20 by thermal induction since the gene is under control of  cI857 repressor

(Cherepanov & Wackernagel, 1995). However, thermal induction of Flp was not desired in our system as this would induce lysogenized P1 mutants as well. It was reported that basal level of Flp induction was sufficient to excise FRT-flanked regions, hence this strategy was applied (Cherepanov & Wackernagel, 1995). Lysogens of P1 mutants, transformed with pCP20 were grown overnight in LB amp + cm at 30 °C. The overnight culture was streaked on LB amp + cm plates to get isolated colonies, from which KanS colonies were selected from patch plating. To ensure that the selected KanS colonies were lysogens of P1 mutants, virions production by thermal induction was tested. The induced P1 mutants were lysogenized into E. coli MG1655 for maintenance of the strains, as previously described and selected as CmR colonies (Piya et al., 2017). The excision of kan gene was further verified by sequencing.

In order to knock out additional genes, P1 mutants were lysogenized into E. coli

BW25113(pKD46) and selected as CmR and AmpR colonies; additional gene was disrupted with FRT-flanked kan gene, as described above.

Efficiency of plating (EOP) assay

All phages used in EOP assays were lysogenized into E. coli WA921 and phages were induced, as previously described (Piya et al., 2017). EOP assays were conducted following previously established protocol, with minor modifications (Piya et al., 2017).

Phages were diluted 10-fold in SM buffer and 10 L from each dilution was spotted on the soft agar overlay of the host strains.

Plasmid construction

The plasmid pBAD24 was engineered to provide gentamicin resistance as described below. The gentamicin resistance cassette was PCR amplified from plasmid pMB838 with primers designed to have AatII and SacI restriction sites on 5’ end of forward and reverse primers, respectively. Similarly, the backbone of plasmid pBAD24 was PCR amplified with primers designed to have SacI and AatII restriction sites on 5’ end of forward and reverse primers, respectively. All PCR reactions were conducted using

Phusion Hi-Fidelity PCR Master Mix (New England Biolabs), following the manufacturer’s recommended protocol. The gentamicin resistance cassette was ligated into pBAD24 backbone using standard molecular biology techniques. P1 genes prt, pro, pmgA, pmgB, pmgC, pmgG and pmgR were cloned into pBAD24g (g for gentamicin resistance). These genes were PCR amplified using primers with XbaI and HindIII restriction sites on 5’ end and were ligated into respective restriction sites in pBAD24g following standard molecular biology techniques.

P1 darB was cloned into pETDuet-1 (Millipore Sigma) for expression and subsequent purification of recombinant DarB by immobilized metal affinity chromatography

(IMAC). The plasmid pETDuet_darB was further engineered to express DarB with 2x

N-terminal His-tag. Synthetic DNA (Integrated DNA Technologies) with regions of

pETDuet-1 vector from lac operator (downstream of T7 transcription start-1) to T7 promoter-2 was obtained, with some modifications. An additional His-tag was introduced downstream of the native His-tag, separated by a linker Gly-Ser-Ser. The introduced His-tag was followed by Ser-Gln to maintain similarity with the native reading frame of pETDuet-1 backbone. Other regions of the pETDuet-1 backbone was not altered. The PCR amplified synthetic DNA fragment was ligated into XbaI and SacI sites in pETDuet_darB following standard molecular biology techniques. To test for the function of recombinant DarB in vivo, DNA fragment encoding DarB with N-terminal

2x His-tag was amplified from pETDuet_2x-His_darB with primers designed to have

XbaI and HindIII restriction sites on 5’ end and ligated into pBAD24 following standard molecular biology techniques.

To determine capsid targeting sequence in DarB, primers were designed with homology to different regions of P1 darB so that several N- and C-terminal residues would be truncated in the expressed mutant DarB. For constructing plasmid expressing DarB with

N-terminal truncations, the start codon ATG was included in the forward primers, whereas for C-terminal truncations, the stop codon TAA was included in the reverse primers. The 5’ end of forward and reverse primers were designed to include XbaI and

HindIII restriction sites. The amplified PCR product was ligated into XbaI and HindIII sites in pBAD24 following standard molecular biology techniques.

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In order to determine if the selected N-terminal residues of DarB were sufficient to target foreign proteins to P1 capsids, two derivatives of plasmid pBAD24 were constructed. Synthetic DNA fragments that encompassed regions of plasmid pBAD24 from 50 bp upstream of NheI restriction site to 50 bp downstream of HindIII restriction site, with some modifications, were ordered. The RBS was modified to “aggaggt”, followed by 8 arbitrary nucleotides. The nucleotides encoding the N-terminal 9 or 30 residues of DarB (including the first residue Met) were added downstream of the arbitrary nucleotides. The pBAD24 multiple cloning sites from EcoRI to HindIII were added downstream of darB specific nucleotides. Both synthetic DNA fragments were

PCR amplified and ligated into NheI and HindIII restriction sites of pBAD24 using standard molecular biology techniques. The two pBAD24 derivatives, pBAD24_darBN9 and pBAD24_darBN30 can be used to clone any genes to express recombinant proteins with selected DarB residues fused to the N-terminus. In this study, tetR-mCherry was

PCR amplified with primers designed to have EcoRI and KpnI restriction sites on the 5’ end and ligated into respective restriction sites of pBAD24_darBN9 and pBAD24_darBN30. Primers and synthetic DNA used in this study are listed in Table

3.2.

Determination of capsid targeting sequence of DarB

Plasmids expressing proteins of interest were transformed into electrocompetent E. coli strain WA921 and transformed cells were selected as AmpR colonies after overnight incubation at 37 °C. The overnight culture of AmpR colonies were lysogenized with

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P1ΔdarB and lysogens were selected on LB amp + cam + kan, as previously described

(Piya et al., 2017). Phage lysates were produced from these P1ΔdarB lysogens as previously described, with minor modifications (Piya et al., 2017). Lysogenic strains were grown at 30 °C in LB amp + cam + kan to OD550 0.4-0.6 and P1ΔdarB was thermally induced at 42 °C. 1 mM L-arabinose (Sigma-Aldrich) was also added at the time of thermal induction. Samples were collected 30 min post-induction and analyzed by SDS-PAGE to check the expression of proteins from the plasmids. The lysates were harvested as described previously (Piya et al., 2017). The phages were purified by CsCl isopycnic centrifugation and the packaging of proteins into capsids were analyzed by

SDS-PAGE of virions.

Purification of virions by CsCl isopycnic centrifugation

Phages were thermally induced in 500 mL LB supplemented with appropriate antibiotics. Phages were concentrated 100-fold by centrifugation and purified by CsCl isopycnic centrifugation as described (Piya et al., 2017).

Size exclusion chromatography of P1 procapsids

P1 mutants disrupted for DNA packaging (P1ΔpacA) were thermally induced from lysogens and lysates were harvested as described before. Protein components were precipitated from lysates by addition of ammonium sulfate to a final saturation of 60%

(Simpson, 2006). The pellet was resuspended in SM buffer, passed through 0.22 m filter (Millipore), followed by size exclusion chromatography on Superdex 200 Increase

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100/300 GL (GE Healthcare). The void volume fractions, comprising P1 proheads and tails, were collected and analyzed by SDS-PAGE.

IMAC protein purification

E. coli BL21(DE3) cells were transformed with plasmid pETDuet_2x-His_darB as previously described (Green & Sambrook, 2012; Piya et al., 2017) with minor modifications. BL21(DE3) cells were concentrated 10x while making them electrocompetent and were transformed with ~2 ng of pETDuet_darB-12His. Cells were recovered in 1mL LB for 1 h at 37 °C and plated to LB amp overnight at 37 °C. Single ampR colony was selected and grown overnight on LB amp. A fresh culture was started in LB amp by 100-fold dilution of the overnight culture. At ~ OD550 0.6-0.8, culture was placed on ice for ~15 mins, 1mM IPTG was added to induce protein synthesis, followed by overnight incubation at 16 °C. Culture was harvested by centrifugation at 5000 x g for 20 mins and washed with cold LB. After additional centrifugation at 5000 x g for 20 mins, cell pellets were stored in -80 °C until further processing.

Cell pellet from 1 L IPTG-induced BL21(DE3) culture was thawed in ice and resuspended in ~30 mL of sonication buffer [50 mM Tris-Cl pH 8.0, 500 mM NaCl,

15% (v/v) glycerol, 10 mM imidazole], followed by addition of 1 mg mL-1 chicken egg white lysozyme (Sigma-Aldrich), 100 g mL-1 bovine pancreas DNase I (Sigma-

Aldrich), SIGMAFAST protease inhibitor (Sigma-Aldrich), 1mM MgSO4 and 0.01%

Triton X-100 (Sigma-Aldrich). The resuspended cells were sonicated to lyse the cells,

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followed by centrifugation for 15 min at 10000 x g at 4°C. The supernatant fraction that contained soluble recombinant DarB was mixed with 5 mL bed volume of pre- equilibrated Ni-NTA agarose resin (Qiagen) in a shaking platform for ~16 h at 4 °C.

DarB bound to Ni-NTA agarose resin was purified by gravity-flow chromatography.

Non-specific proteins bound to resin was eluted with IMAC wash buffer (50 mM

Sodium phosphate pH 7.4, 500 mM NaCl) with gradient concentration of imidazole (20 mM, 50 mM, 60 mM, 70 mM, 80 mM, 100 mM). The target protein was eluted with elution buffer (50 mM Sodium phosphate pH 7.5, 500 mM NaCl, 500 mM imidazole).

The elution fraction was concentrated, and buffer exchanged (50 mM Sodium phosphate pH 7.4, 150 mM NaCl) in 50 kDa molecular weight cutoff centrifugal filters (Amicon).

The concentrated protein was stored in 20% glycerol (v/v) at -20°C until further use

(Green & Sambrook, 2012). Samples were collected at each step for SDS-PAGE analysis.

SDS-PAGE

Phage samples for SDS-PAGE analysis were prepared as previously described (Piya et al., 2017). For other protein samples, DNase I treatment was omitted. The gels were stained with either SYPRO Ruby (Thermo Scientific) or Coomassie (GE Healthcare).

Transmission electron microscopy

Samples for electron microscopy were stained with 2% uranyl acetate and imaged in a

JEOL 1200 EX transmission microscope under 100 kV accelerating voltage in the

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Microscopy and Imaging Center, Texas A&M University, College Station (Valentine et al., 1968). Crude lysates of P1 procapsids were concentrated 10-fold in 50 kDa MWCO centrifugal filters (Amicon) for imaging.

Results and discussion

P1 genes of unknown function are not involved in antirestriction system

The first reported antirestriction phenotype of phage P1 demonstrated that two P1 genes darA and darB, comprised the P1 antirestriction system (Iida et al., 1987). It has been shown that darA and darB are present in two distinct operons in P1 genome (Lobocka et al., 2004). Recent work has shown that along with darA and darB, other genes present in the operons encoding these two genes, hdf, ddrA, ddrB and ulx are associated with the

P1 antirestriction system (Fig. 3.1) (Piya et al., 2017). Moreover, hdf and darA have been depicted to be involved in determination of head size during virion morphogenesis

(Piya et al., 2017).

Since the genes encoding P1 antirestriction system were present in two different operons in P1 genome, it was hypothesized that genes elsewhere in the P1 genome could be involved in providing protection against type I R-M systems. The identity of the antirestriction protein protecting P1 DNA from the type I EcoA system is still not clear.

Moreover, proteins Hdf and DarA from the P1 antirestriction system were also found to affect capsid morphogenesis (Piya et al., 2017). P1 capsid morphogenesis has not been well studied and the proteins forming the scaffolding core have not been identified

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(Lobocka et al., 2004; Piya et al., 2017). Despite the role of Hdf and DarA in determining normal P1 capsid size, they do not fit the classical definition of scaffolding proteins, as both Hdf and DarA are still present in the mature virions (Piya et al., 2017).

We attempted to study the role of other P1 genes in antirestriction system and virion morphogenesis. There are several genes in P1 genomes for which any definitive function has not been assigned. The genes that were suggested to have unknown function or putative morphogenetic functions were of interest to us (Lobocka et al., 2004). Isogenic deletions of 29 such genes in P1 genome were made and the role of each gene was interrogated in relation to P1 antirestriction system and virion morphogenesis (Fig. 3.1).

Successful isogenic deletions of these 29 genes in P1 lysogens suggest that these genes do not affect stability of P1 lysogens. P1 mutants deleted for genes pmgA, pmgB, pmgC, pmgG or pmgR did not form plaques on lawns of E. coli host, suggesting these genes are essential for lifecycle of P1. The essentiality of these five genes has not been verified by in trans complementation.

The remaining 24 gene knockout mutants of P1 were able to form plaques on lawns of E. coli host. These P1 mutants were lysogenized into E. coli WA921 strain and phage lysogens were induced to check for restriction phenotype. It has been established that the

EOP of P1ΔddrA is severely reduced in cells with type I EcoA, EcoB or EcoK systems as virions P1ΔddrA mutants are unable to incorporate antirestriction proteins in capsids during virion morphogenesis (Piya et al., 2017). The EOP of all P1 mutants were

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compared to the EOP of P1ΔddrA to study roles played by those genes in antirestriction.

In this study, the EOP of P1ΔddrA was reduced by ~10-3, 10-4 or 10-4 in E. coli strains with type I EcoA, EcoB or EcoK systems respectively. The EOP of all other phage mutants appeared ~1 in E. coli strain with type I EcoA system and ~10-1 - 10-2 in E. coli strains with type I EcoB or EcoK systems (Fig. 3.2). This suggests that the antirestriction system of these mutants were not compromised and these phage mutants were able to employ antirestriction system to protect DNA from type I R-M system mediated DNA cleavage following infection of the respective host.

Dar proteins are packaged into P1 capsids before DNA

The Dar antirestriction system of P1 is different from other studied antirestriction systems as Dar proteins act strictly in cis (Iida et al., 1987). These Dar proteins are incorporated into capsids during virion morphogenesis and at least some proteins must be delivered into the host cytoplasm during phage infection, where they protect DNA from host restriction (Iida et al., 1987; Piya et al., 2017). Recent work has revealed that

Dar antirestriction proteins are incorporated into P1 capsids in a specific order during virion morphogenesis (Piya et al., 2017). The assembly pathway of most phages in

Caudovirales order bear similarity in that assembly starts with prohead, which is eventually filled with genetic material (Aksyuk & Rossmann, 2011). Phage P1 is not unique in packaging proteins into the head which are later transferred to the host cytoplasm on infection. Phage T4 also has several internal proteins packaged into procapsids during capsid morphogenesis. These internal proteins of T4 are incorporated

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into scaffolding core, followed by procapsid shell formation and DNA packaging (Hong

& Black, 1993; Leiman et al., 2003). In double-stranded DNA phages, DNA is packaged

Fig. 3.2. P1 genes of unknown function are not required for antirestriction. Phage P1 or mutants were lysogenized into restriction-modification deficient E. coli strain WA921. The lysogens were thermally induced and tested for plating efficiency in E. coli strains with type I EcoA, EcoB or EcoK R-M systems. The EOP of parental P1, -1 -3 -2 with functional antirestriction system is ~10 , ~10 or 10 in strains with type I EcoA, EcoB or EcoK systems respectively, whereas the EOP of P1ΔddrA, with -3 -4 -4 -5 disrupted antirestriction system, is ~10 , ~10 or ~10 -10 in strains with type I EcoA, EcoB or EcoK systems respectively. The EOP of all other P1 mutants generated in this study appeared normal compared to the EOP of P1ΔddrA, suggesting that these genes do not play any roles in protecting P1 DNA from the type I R-M systems tested in this study.

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Fig. 3.2. Continued.

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into procapsids by a packaging complex, composed of small (TerS) and large (TerL) subunits (Aksyuk & Rossmann, 2011). The DNA packaging complex of P1, which is composed of proteins encoded by pacA and pacB, cleave P1 DNA concatamers at pac sites located within the coding region of pacA (Fig. 3.3A) (Skorupski et al., 1992). In the absence of either PacA or PacB, DNA cannot be packaged into P1 procapsids and virion morphogenesis is stalled at the procapsid stage (Skorupski et al., 1992).

In order to determine if the antirestriction proteins are incorporated into P1 procapsids before DNA, pacA was deleted from P1 prophage. These ΔpacA prophages were then induced and procapsids were purified by ammonium sulfate precipitation followed by size exclusion chromatography as described above in materials and methods. These methods are suitable for purification of macromolecular complexes, hence both procapsids and tails are purified in these fractions. These procapsid/tail fractions as well as CsCl purified intact parental P1 virions were then analyzed by SDS-PAGE (Fig.

3.3B). Previously identified virion-associated Dar proteins, DarB, DarA, DdrB and Hdf, have been labelled. Other proteins, Ulx and DdrA, which are also associated with the

Dar antirestriction system are not seen in this SDS-PAGE because of their small size and low copy number. It has been established that DarB can be incorporated into virions efficiently only in the presence of all other Dar proteins. Thus, the presence of DarB is indicative of the presence of other Dar proteins in the capsids. In the lanes of both CsCl

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Fig. 3.3. P1 antirestriction components are incorporated into P1 procapsids before DNA. Panel A: The image represents a partial genomic map of P1 showing genes pacA and pacB, required for DNA packaging during virion morphogenesis. Both pacA and pacB are transcribed rightwards. The ruler at the bottom represents the nucleotide position as in the published P1 genome (NC_005856.1). The gene pacA was deleted from P1 genome in this study. Panel B: Large macromolecular structures present in thermally induced lysates of P1ΔpacA and P1ΔdarA_ΔpacA were concentrated by ammonium sulfate precipitation and fractionated by size exclusion chromatography. The void volume from size exclusion chromatography, consisting of P1 procapsids and tails, was collected and analyzed by 12% Tris-glycine SDS-PAGE. P1 virions, purified by cesium chloride isopycnic centrifugation, were also run in SDS-PAGE for comparison. The protein bands corresponding to the components of P1 antirestriction system seen in the SDS-PAGE have been annotated. Presence of proteins bands corresponding to DarB, DarA, DdrB and Hdf in the lane of P1ΔpacA indicates that these antirestriction components are packaged into P1 procapsids before DNA. Other protein bands appearing in the procapsid lanes are denoted by arrows.

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Fig. 3.3. Continued.

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purified intact P1 virions and P1ΔpacA fractions, the band corresponding to DarB is visible. However, it is possible that this visible protein band of DarB in P1ΔpacA lane is not associated with the procapsids, but co-purified from the lysates at the applied ammonium sulfate concentrations (Wingfield, 2001). In order to rule out this possibility, procapsid/tail components of P1ΔdarA_ΔpacA were also purified from crude lysates and were analyzed simultaneously (Fig. 3.3B). P1ΔdarA virions are unable to package DarB, thus it is expected that P1ΔdarA_ΔpacA procapsids also cannot package DarB (Piya et al., 2017). However, the darB gene is intact in the genome of the P1ΔdarA prophage, thus DarB is synthesized normally when lysogens of P1ΔdarA are thermally induced, despite DarB not being incorporated into capsids (Fig. 3.4). A protein band corresponding to DarB is not visible in the lane of P1ΔdarA_ΔpacA, indicating that

DarB proteins are not incorporated into procapsids, and also are not purified along with procapsid/tail fractions by this technique despite being present in lysates. As mentioned above, in the absence of PacA, virion morphogenesis is stalled at the procapsid stage as the DNA packaging event is disrupted. Hence, comparative analysis of protein bands of

P1ΔpacA and P1ΔdarA_ΔpacA mutants suggest that DarB and thus other antirestriction proteins are packaged into procapsids before DNA.

Electron densities in transmission electron micrographs of P1 procapsids suggest presence of other internal proteins

In negative-stained transmission electron micrographs (TEM) of P1ΔpacA lysates, empty procapsid structures were visible (Fig. 3.5). It was interesting to observe internal

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Fig. 3.4. DarB is synthesized in P1ΔdarA mutants. E. coli MG1655 lysogens of P1

and P1ΔdarA were cultured in 30 °C up to OD550 0.4-0.5 and thermally induced at 42 °C. Samples were collected before and at 15, and 30 min post-thermal induction. 0.3 OD equivalent of samples were loaded in 8% Tris-glycine SDS-PAGE and gel was stained with SyproRuby.

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A B

C D

Fig. 3.5. P1 procapsids show electron densities suggesting presence of protein contents. Images shown are representative negative-stained transmission electron micrographs of procapsids of P1 (A), P1ΔdarA (B and C) and P1ΔdarB (D), generated by deletion of DNA packaging component pacA from P1 genome. Since these mutants are defective in DNA packaging, the presence of electron density can be corroborated to presence of proteins. Presence of electron density in P1ΔdarA (B and C) and P1ΔdarB (D) indicate these densities correspond to proteins other than Dar proteins. These electron densities are also seen in both normal (B) and small (C) size class procapsids of P1ΔdarA. The scale bar represents 100 nm.

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electron densities inside these protein shells. In a complete virion, this electron density inside capsid suggests packaged DNA (Chapter II, Fig 2.8). However, P1ΔpacA is defective in DNA packaging, thus it is extremely unlikely that this density is from the packaged DNA. It has been demonstrated that P1 Dar proteins are localized into P1 capsids and contribute significantly to the mass of the virion (Iida et al., 1998; Piya et al., 2017). Hence, we speculated that these densities correspond to the internal Dar proteins. However, the electron densities were still conspicuous in the negative-stained micrographs of both P1ΔdarB_ΔpacA (Fig. 3.5D) and P1ΔdarA_ΔpacA (Fig. 3.5B and

3.5C). P1ΔdarB virions are missing DarB and Ulx, whereas P1ΔdarA virions are missing all known proteins of P1 Dar system (Piya et al., 2017). Thus, it can be concluded, from SDS-PAGE and TEM analysis of P1ΔdarA_ΔpacA procapsids, that the electron densities visible in P1 procapsids do not correspond to the antirestriction proteins. Negative-stained TEM images of phage P22 and T4 procapsids also show similar electron density inside the capsid shells that correspond to scaffolding protein core (King et al., 1973; Traub & Maeder, 1984; Black & Rao, 2012). Thus, it is possible that the density observed in P1 procapsids also correspond to the scaffolding core, which might be removed during DNA packaging (Fig. 3.5).

Comparative SDS-PAGE analysis protein bands of P1ΔpacA and P1ΔdarA_ΔpacA show that the lanes of these procapsids contain protein bands (marked with arrows) that are absent in the lanes of CsCl purified intact P1 virions (Fig. 3.3B). Procapsids of other phages such as P22 and T4 contain scaffolding proteins that are either recycled or

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cleaved and are not present in mature virions (Dokland, 1999). These protein bands in

P1 procapsid lanes could be the constituents of the P1 scaffolding core. Because of the limitations of the technique applied in this study to obtain procapsids, it is also possible that some protein bands in the procapsids lanes of Fig. 3.3B are contaminating host proteins, thus the identity of these proteins need to be confirmed by mass spectrometry.

N-terminal residues of DarB provides signal for capsid targeting

Phages such as T4 and P22 incorporate internal proteins into capsids during early stages of capsid morphogenesis (Hong & Black, 1993; Jin et al., 2015). The capsid-associated phage T4 internal protein III (IPIII) is directed to the assembly core by specific N- terminal residues, designated as capsid targeting sequence (CTS) (Showe & Black, 1973;

Mullaney & Black, 1996). Since DarB is also packaged into the P1 capsid, it is possible that DarB has a CTS that directs nascent DarB to P1 procapsids. To determine if DarB is directed to capsid assembly by any specific residues, mutant DarB truncated at the N- and/or C-terminus were expressed from a plasmid-based system. Full-length DarB consists of 2255 residues, with a mass of ~250 kDa. Initially, three versions of truncations were constructed: a C-terminal truncation, with DarB containing residues 1-

1750 (DarBt 1-1750), an N-terminal truncation, with DarB containing residues 90-2255

(DarBt 90-2255) and both N- and C-terminal truncations, with DarB containing residues

90-1750 (DarBt 90-1750). The truncated DarB was induced in P1ΔdarB lysogen and the virions were tested for restoration of antirestriction phenotype and for incorporation of

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truncated DarB. Two conserved domains of DarB were kept intact while constructing all truncated versions.

EOP assays shows that none of these truncated DarB restored the antirestriction phenotype of P1ΔdarB mutant (Fig. 3.6A). In order to determine if the truncated DarB was expressed, whole cell samples of induced lysogens were analyzed via SDS-PAGE

(Fig. 3.7A). DarB expressed from parental P1 is not visible because of the sensitivity of the staining procedure. However, the presence of DarB in P1virions is evident from the normal EOP of P1 in strains with type I EcoB or EcoK (data not shown). Protein bands corresponding to expressed protein were visible for all truncations suggesting that DarB truncations were expressed at a higher level from plasmid-based system compared to wt

DarB expressed from its native locus in P1.

To determine if any of these DarB truncations were packaged into virions, P1ΔdarB virions complemented in trans with DarB truncations were purified by CsCl isopycnic gradient centrifugation and analyzed by SDS-PAGE. Of the three truncations, protein bands corresponding to only DarBt 1-1750 was visible in the SDS-PAGE of virions.

Protein bands corresponding to two other truncations, DarBt 90-1750 and DarBt 90-

2255, were not visible. This suggests the N-terminal ~89 residues of DarB are required for targeting DarB into capsid during capsid morphogenesis (Fig. 3.8A). However, since

DarBt 1-1750 cannot restore the antirestriction phenotype of P1ΔdarB virions, it is

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EOP

Fig. 3.6. Plating efficiency of P1ΔdarB complemented with truncated DarB. In order to test for packaging signal in DarB, several N- and C-terminal truncations of DarB were constructed and tested for their ability to complement restriction phenotype. The empty vector pBAD24 or pBAD24 cloned with darB fragments were transformed into E. coli strain WA921 and lysogenized with P1ΔdarB. The phage lysates were obtained as described in materials and methods and were tested for restoration of plating efficiency on E. coli strains with type I EcoA, EcoB or EcoK R- M systems. The EOP data shown have been normalized to P1 induced from strain WA921 transformed with plasmid pBAD24. Panel A: The relative EOP of P1ΔdarB -2 -3 was reduced by ~10 and 10 in EcoB and EcoK strains respectively (red bars). The EOP was restored to normal levels by full length DarB (green bars). None of the truncated DarB was able to restore the EOP (grey bars). Panel B: As described in panel A, the EOP of P1ΔdarB was reduced in strains with EcoB and EcoK strains (red bar) and the EOP was restored to normal levels by full length DarB (dark green bar). N-terminal truncation of DarB containing residues 5-2255 was able to restore plating deficiency in EcoB and EcoK strains (light green bar). None of the other truncations restored plating deficiency in EcoB and EcoK strains (grey bars). In both panels A and B, the plating efficiency was not affected in EcoA strain. The data shown are averages of two biological replicates.

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EOP

Fig. 3.6. Continued.

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Fig. 3.7. Truncated DarB can be expressed. For both panels A and B, E. coli strain WA921 was transformed with empty plasmid pBAD24 or plasmid encoding full length or truncated DarB as denoted. Transformed cells were lysogenized with P1 or P1ΔdarB as indicated. Phages and protein production from plasmids were induced simultaneously, as described before. Samples were collected ~30 min post-induction and analyzed by SDS-PAGE and Coomassie staining to check for protein induction from plasmids. Both full-length and truncated versions of DarB are induced from the plasmids, as evident from presence of corresponding protein bands.

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Fig. 3.8. N-terminal residues of DarB provide signal for capsid targeting. For both panels A and B, phages P1 or P1ΔdarB were lysogenized into E. coli strain WA921 containing respective plasmid. Phages and proteins from plasmids were induced simultaneously as described before. Phages were purified by cesium chloride isopycnic gradient centrifugation and analyzed by SDS-PAGE, followed by SyproRuby staining.

Panel A: DarBt 1-1750 is incorporated into P1ΔdarB virions. Panel B: DarBt 5-2255 is packaged into P1ΔdarB virions. All other truncations of DarB are not packaged into P1ΔdarB virions, suggesting few N-terminal DarB residues provide signal for capsid targeting.

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Fig. 3.8. Continued.

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uncertain if DarBt 1-1750 can be ejected into host cells during infection. Moreover,

DarBt 1-1750 is missing ~500 C-terminal residues. Even though there is no identifiable conserved domain within these C-terminal 500 residues of DarB, these 500 residues might be necessary for folding of DarB into native conformation or for establishing other protein-protein or protein-DNA interactions to perform its functions.

Once we established that N-terminal residues of DarB are required for capsid localization, we sought to determine the specific residues required for this purpose.

Several constructs were produced to express truncated DarB translated from specific N- terminal residues: 5 (DarBt 5-2255), 10 (DarBt 10-2255), 15 (DarBt 15-2255), 20 (DarBt

20-2255) and 25 (DarBt 25-2255). Of these truncations, only DarBt 5-2255 was able to complement the antirestriction phenotype of P1ΔdarB (Fig. 3.6B). SDS-PAGE of the induced lysogens showed that other four truncated DarB proteins were expressed from plasmids as well (Fig. 3.7B). P1ΔdarB virions complemented with these series of truncated DarB were purified as described above and analyzed by SDS-PAGE. Protein band corresponding to DarBt 5-2255 was visible in SDS-PAGE, which corelates to why

DarBt 5-2255 was able to complement antirestriction phenotype of P1ΔdarB. Neither of the truncated DarB starting with residues 10, 15, 20, and 25 were packaged into the virions (Fig. 3.8B). This suggests that N-terminal amino acid sequences of DarB spanning residues 5 to 9 are crucial for targeting DarB into procapsids during capsid morphogenesis.

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Purification of DarB

Genetic evidence suggests that DarB protects P1 DNA from host type I R-M system mediated DNA cleavage, following infection (Iida et al., 1987; Piya et al., 2017).

However, biochemical mechanisms of DNA protection have not been elucidated. More importantly, it is not known if DarB needs any interacting partners to perform its biological role. Bioinformatic analysis shows that P1 DarB consists of a methyltransferase and a DExH helicase domain (Gill et al., 2011). Based on the presence of conserved domains, biochemical activity of DarB can be predicted, however this must be confirmed in vitro. Since much of this future work will depend on the ability to obtain purified DarB protein, we sought to purify DarB by using affinity chromatography. Optimization of purification steps would also facilitate the study of the interacting partners of DarB by conducting pull-down assays (Green & Sambrook,

2012).

Initial attempts were made to purify DarB by cloning P1 darB into expression vector pETDuet-1 in-frame with the native His-tag. However, eluted DarB was not of high purity. Other available tags, glutathione S-transferase (GST) (GE Healthcare), maltose- binding protein (MBP) (New England Biolabs) and chitin binding domain (CBD) (New

England Biolabs) were explored by fusing them to the N-terminus of DarB. Fusion of these tags to DarB produced varying results: inefficient binding of recombinant DarB to purification resin (GST-tag), toxicity (MBP-tag) and no protein expression (CBD-tag).

Because of the relatively large size of DarB (~250 kDa), we reasoned that a single His-

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tag did not provide sufficient binding affinity to the recombinant DarB. Hence, an additional His-tag was introduced into the backbone of pETDuet-1, downstream of the native His-tag, resulting in the production of recombinant DarB with 2x His-tag in its N- terminus (2x-His_DarB). The recombinant DarB has the following residues on its N- terminus: MGSSHHHHHHGSSHHHHHHSQDPNSSS (the underlined residues indicate

His-tag)

Affinity purification of recombinant 2x-His_DarB yielded protein with significant purity as determined by SDS-PAGE analysis (Fig. 3.9). Even though His-tags are smaller compared to other affinity tags, they can still affect biochemical properties of recombinant proteins (Booth et al., 2018). Thus, we sought to determine if 2x-His_DarB can complement the restriction phenotype of P1ΔdarB. The gene encoding 2x-His_DarB was PCR amplified and cloned into pBAD24 vector to test for function of 2x-His_DarB in vivo. EOP assays suggest that 2x-His_DarB can restore the restriction phenotype of

P1ΔdarB (Fig. 3.10). Thus, it can be inferred that 2x His-tag fused to N-terminus of

DarB does not affect the biochemical activity of DarB significantly, at least in vivo.

Conclusions

In Chapter 2 of this dissertation, it was demonstrated that P1 antirestriction is comprised of multiple components, in addition to the established darA and darB (Iida et al., 1987;

Piya et al., 2017). Moreover, the role of hdf and darA in head size determination during virion morphogenesis suggest that P1 antirestriction is also linked to capsid

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Imidazole step-gradient wash

250 2x-His_DarB 150

100

Imidazole step-gradient wash

250 2x-His_DarB 150 100

Fig. 3.9. IMAC purification of P1 DarB. E. coli BL21(DE3) strain was transformed with the expression plasmid pETDuet_2x-His_darB and protein production was induced by addition of 1 mM IPTG. BL21(DE3) cells expressing 2x His_DarB were sonicated and the supernatant (Sup) was applied to Ni-NTA agarose resin. Flow-through (FT) fraction was collected and non-specific proteins bound to the resin was washed with buffer containing increasing gradient of imidazole. 2x His_DarB bound to Ni-NTA resin was eluted with buffer containing 500 mM imidazole. All fractions were collected and analyzed by SDS-PAGE, followed by Coomassie staining.

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Fig. 3.10. DarB with 2x His-tag fused to N-terminus is functional. Phages for complementation were induced as described before. The EOP of P1ΔdarB is reduced in E. coli strains with EcoB or EcoK systems, as demonstrated before (red bar). DarB with 2x His-tag fused to its N-terminus can restore the plating defect of P1ΔdarB in strains with EcoB or EcoK systems suggesting that the N-terminal fusion of 2x His- tag does not disrupt function of DarB (green bar). The data shown are averages of two biological repeats.

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morphogenesis. As the components of the P1 Dar system were distributed in two operons in P1 genome (Piya et al., 2017), we sought to interrogate genes in other operons for their role in antirestriction and/or capsid morphogenesis. We targeted P1 genes of unknown function for this study. Isogenic deletion knockouts of these genes were created in P1 lysogens and induced virions were tested for restriction phenotype in

E. coli hosts with type I EcoA, EcoB or EcoK systems by conducting plating assays

(Fig. 3.1). The efficiency of plating (EOP) of all of these P1 mutants were comparable to the EOP of parental P1, which suggests that the antirestriction system is intact in these mutants (Fig. 3.2). Thus, these P1 genes do not appear to contribute to the activity of the

P1 antirestriction system.

Lysates of knockout mutants of five P1 genes pmgA, pmgB, pmgC, pmgG and pmgR did not form plaques on lawns of their E. coli host (Fig. 3.1), indicating that these five genes are essential for plaque formation under laboratory conditions. At this stage, it is too early to ascribe any functions to these genes as plaque non-forming phenotype of these mutants still needs to be complemented. Genes pmgA, pmgB, pmgC and pmgG are found in vicinity of other genes encoding tail components. Although it is tempting to speculate that these genes encode components of tail assembly based on their location in P1 genome, it might be misleading to do so as genes encoding components of same pathway are not organized together in P1 genome. After complementing the phenotype pertaining to those genes, lysates of mutant virions may be analyzed by transmission electron microscopy to ascertain potential effects of these genes on virion morphogenesis.

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Virions such as P22 and T4 incorporate several internal proteins into procapsids during earlier stages of capsid morphogenesis, followed by DNA packaging (Bazinet & King,

1988; Arisaka, 2005; Aksyuk & Rossmann, 2011). By disrupting DNA packaging in P1, virion morphogenesis was stalled at formation of procapsids and tail and both procapsids and tails were purified in same fraction. Comparative SDS-PAGE analysis of the procapsids/tail fractions of P1ΔpacA and P1ΔdarA_ΔpacA suggest that P1 antirestriction components are packaged before DNA (Fig. 3.3).

P1 Dar proteins are incorporated in capsids and at least one of these, DarB, is injected into the new host during infection. As phage DNA ejected into host are packaged in the inner space of capsids, it can be extrapolated that these Dar proteins are also packaged in the same space. It was demonstrated in Chapter 2 of this dissertation that P1 Dar proteins occupy a significant mass in the capsid (Piya et al., 2017), which suggests that not having Dar proteins inside of P1 procapsids will leave more space for DNA to be packaged. Virions of P1ΔdarA cannot package any of the Dar proteins, thus internal space in P1ΔdarA procapsids is likely to be greater than those of P1 procapsids. Since P1 packages DNA into procapsids by headful mechanism (Bachi & Arber, 1977; Lobocka et al., 2004), virions of P1ΔdarA should be able to package more DNA than P1, which can be tested by pulse-field gel electrophoresis of genomic DNA (Lingohr et al., 2009).

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The internal volume of P1 procapsids, as seen in negative-stained EM images, has an asymmetrically distributed electron density, suggestive of proteins because P1 procapsids constructed in this study are unable to package DNA (Fig. 3.5). Procapsids harvested at similar stages of virion morphogenesis in other phages such as P22 and T4 also show similar electron densities, which are attributed to the scaffolding core.

However, this scaffolding core of both P22 and T4 occupy most of the space inside of procapsid, compared to partial space in P1 (King et al., 1973; Traub & Maeder, 1984).

This suggest that the scaffolding core of P1 is either proteolytically processed and the fragments are removed before the start of DNA packaging or there is a novel mechanism associated with head morphogenesis. Identification of proteolytic processing enzyme and scaffolding proteins associated with P1 head morphogenesis, if there is any, would assist in studying P1 capsid morphogenesis.

Comparative SDS-PAGE analysis of CsCl purified mature P1 virions and P1 procapsids/tail fractions show few proteins bands that are only present in P1 procapsids/tail fractions. These new protein bands present in the lanes of procapsids/tail fractions are suggestive of constituents of scaffolding core, as this core is processed and removed during later stages of capsid morphogenesis (Dokland, 1999). However, due to the limitations of current purification methods, P1 procapsids and tail components are purified in same fractions. Moreover, these fractions may contain contaminating host proteins. After optimization of procapsids purification, it can be determined from SDS-

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PAGE if these proteins bands are still present in the lanes of procapsids and can be identified by mass spectrometry.

P1 antirestriction protein DarB is incorporated into capsid in a definite pathway involving other capsid-associated proteins as well (Piya et al., 2017). By analyzing incorporation of DarB mutants, truncated for several residues at N- or C- terminus, into

P1ΔdarB virions, we have determined that N-terminal residues ranging from 5th to 9th are essential for packaging of DarB into P1 capsids (Fig. 3.8B). Residues that are crucial for targeting proteins into capsids have also been found in other phages. Phage T4 capsids contain hundreds of copies of internal proteins (IPI, IPII and IPIII) (Black &

Ahmad-Zadeh, 1971). It has been determined that IPIII is incorporated into T4 procapsids with a capsid targeting sequence (CTS) present in N-terminus of this protein

(Mullaney & Black, 1996). Moreover, the CTS can also direct other foreign proteins to be packaged into T4 capsids and these foreign proteins are further injected into new host during T4 infection (Hong & Black, 1993; Mullaney & Black, 1996). Although DarB N- terminal residues 5th-9th seem to be required for packaging of DarB into P1 virions, it is not evident if those residues are sufficient. Fusion of foreign proteins to the respective

N-terminal residues of DarB could tell if these particular residues are sufficient for packaging any protein into P1 capsids. Moreover, fusion of fluorescent proteins could be advantageous as injection of these fusion proteins into new host following P1 infection can be tested using microscopy.

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Although it has been established that DarB protects P1 DNA from type I R-M system mediated DNA cleavage following ejection of DNA into host cytoplasm (Iida et al.,

1987; Piya et al., 2017), biochemical mechanism behind this protection has not been determined. Because of the presence of conserved methyltransferase and helicase domains in DarB, it may be hypothesized that DarB methylates and/or translocates along

DNA following ejection of both DarB and DNA into new host (Lobocka et al., 2004;

Gill et al., 2011). In order to determine the activity of DarB in vitro, we sought to purify

DarB using affinity chromatography. By fusion of 2x His-tag to N-terminus of P1 DarB, recombinant DarB could be purified, by immobilized metal affinity chromatography, to sufficient purity to conduct enzyme assays (Fig. 3.9). Moreover, we have also shown that 2x His-tag fusion to DarB does not abolish its biological activity (Fig. 3.10). In vitro enzyme assays can now be conducted to determine the biochemical mechanism of how

DarB protects DNA from type I R-M system mediated restriction.

Many proteins can perform biological functions only when they are associated with other protein subunits forming a complex. Thus, it cannot be ruled out yet that DarB could only function in presence of other protein subunits. In the type I R-M system complex, DNA specificity is provided by HsdS subunit (Murray, 2000). Since DarB has been shown to protect DNA against other type I R-M systems from Salmonella as well, it is plausible that DarB might interact with HsdS subunit of type I R-M complex (Iida et al., 1987). Moreover, other proteins from Dar system might be co-injected with DarB during phage infection. Since affinity tag-based purification of DarB has been optimized,

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same strategy can be applied to set up pull-down assays to determine interacting partners of DarB. Setting up in vitro enzyme assays for DarB may be worthwhile, only after discovering interacting partners of DarB, if any.

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CHAPTER IV

GENOME-WIDE SCREENS REVEAL ESCHERICHIA COLI GENES

REQUIRED FOR GROWTH OF T1-LIKE PHAGE LL5 AND RV5-LIKE

PHAGE LL12

Introduction

Escherichia coli was discovered by the German microbiologist Theodor Escherich, in the study of infant gut microbes (Escherich, 1988). Because of several traits such as easily culturable, short doubling time, E. coli has been used as a model organism for understanding basic biology as well as for biotechnology applications. E. coli is a member of Enterobacteriaceae, and is related to other pathogens such as Salmonella,

Klebsiella, Serratia and Yersinia. E. coli is a facultative aerobe, and along with other obligate anaerobes, forms a part of commensal gut flora (Blount, 2015). These commensal E. coli strains do not cause disease in human host, barring immunocompromised individuals. However, there are several strains of E. coli that have acquired virulence factors and cause diseases with symptoms, ranging from mild discomfort to life-threatening. Depending upon disease etiology, these strains have been categorized into several pathotypes: enteropathogenic E. coli (EPEC), enterohemorrhagic E. coli (EHEC), enterotoxigenic E. coli (ETEC), enteroaggregative

E. coli (EAEC), enteroinvasive E. coli (EIEC), and diffusely adherent E. coli (DAEC)

(Kaper et al., 2004).

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This study aims to study phage-host interactions between E. coli and two virulent phages, LL5 and LL12, which were isolated against ETEC host strains. ETEC strains can be distinguished from other E. coli pathotypes by the presence of two toxins: heat- labile enterotoxins (LTs) and heat-stable enterotoxins (HTs). These strains might express only one or both of those toxins (Kaper et al., 2004). These enterotoxins induce traveler’s diarrhea (TD), characterized by watery diarrhea, for which the symptoms could range from mild to severe (Kaper et al., 2004). TD is one of the most common illnesses contracted by people from developed countries travelling to less developed countries of the world. TD is also accompanied by other symptoms such as nausea, vomiting, abdominal pain, fever or blood in stool (Taylor et al., 2017). The causative agent of TD varies in different geographical regions; Campylobacter spp. is the primary one in Southeast Asia whereas ETEC is the major one in the Latin America, Africa, south Asia and the Middle East (Tribble, 2017). Besides, other bacterial pathogens such as EAEC, Shigella spp., non-typhoidal Salmonella spp., and viruses and parasites have also been reported to cause TD (Taylor et al., 2017; Tribble, 2017). TD has been, so far, successfully treated with antibiotics, but the global increase in the emergence of the antibiotic resistance in bacteria warrants evaluation of alternative treatment approaches

(Tribble, 2017).

Bacteriophages (phages) are the natural predators of bacteria. Due to emergence of multidrug resistant pathogens and limitation in the discovery of antibiotics, there has been a renewed interest in phage therapy in western medicine. Phage therapy is the

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application of phages to treat pathogenic bacteria (Young & Gill, 2015). Phage therapy could be a solution to increasing antibiotic resistance incidence in the causative agents of

TD. Phage infection cycle starts with adsorption of phages to host receptors, ejection of phage genetic material into host, replication of genetic material and production of phage structural components, followed by lysis of host cells to liberate progeny phages to surroundings. As bacteria are constantly preyed upon by these phages, bacteria are evolved with diverse defense mechanisms to thwart phage infection (Labrie et al., 2010).

Hence, for phage therapy to succeed, the necessity of thorough understanding of phage- host interactions cannot be overstated.

Phage replication needs to be very robust because phage infection cycle is short. Phages may not carry all genes required for necessary functions because there is a limitation in their genome size. Therefore, to have an efficient replication cycle, phages utilize various host functions. For instance, phages need quick and abundant chaperone power so that the assembly of virions can be completed before host lysis (Georgopoulos, 2006).

Same holds true for eukaryotic viruses as well. Before the advent of modern genetic tools, phage-host interactions were studied using classic genetic tools. Despite these classic approaches being powerful, it does have its limitations as these techniques are labor-intensive and lengthy. With the availability of modern genetic resources, the large- scale host-phage interactions can be studied efficiently. By applying these modern genetic approaches, a number of genome-wide screens have been conducted to study host factors required for viral replication in organisms such as HIV (Brass et al., 2008),

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Influenza virus (Hao et al., 2008), phage λ (Maynard et al., 2010) and phage T7 (Qimron et al., 2006).

We conducted forward-genetics screen of phages, LL5 and LL12 against the Keio collection to characterize any host function that is required for efficient phage replication. The Keio collection is a library of single-gene deletions of all non-essential genes in E. coli K-12 strain BW25113 (Baba et al., 2006). This screen will help in understanding host-virus interaction (Maynard et al., 2010) by revealing host factors involved in infection and replication of LL5 and LL12. The phages will not be able to infect and propagate efficiently in a host devoid of certain crucial functions, resulting in these mutant hosts outgrowing the WT host, upon simultaneous infection of the phages.

Identification of host factors required for phage propagation is crucial for the development of therapeutic phages because the bacterial host can develop resistance to phages by mutating the host factors, resulting in absolute protection from phage infection and propagation. If the knowledge about host factors is available, then the phage genome can be engineered to include those factors so that there is reduced dependency in the host genome encoded factors (Qimron et al., 2006). More importantly, if the host receptors can be characterized for individual phages, phage cocktails, composed of phages targeting different host receptors, can be prepared, which will improve the efficacy of phage therapy. Understanding host-virus interaction is very crucial for the application of phage therapy because it will facilitate the engineering of

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robust phages (Qimron et al., 2006). The results of this screen are discussed in terms of host factors required for infection and propagation of phages LL5 and LL12.

Materials and methods

Bacterial strains and plasmids

The Keio collection was purchased from Thermo Scientific (Baba et al., 2006). Strains from the ASKA library used for complementation were purchased from National

BioResource Project (NIG, Japan) (Kitagawa et al., 2005). The parental E. coli strain

BW25113 was obtained from Ry Young (Texas A&M University, College Station, TX).

E. coli strains from the Keio collection and their transductants were cultured in LB

(Lennox) broth [10 g L-1 Bacto tryptone (BD), 5 g L-1 Bacto yeast extract (BD), 5 g L-1

NaCl (Avantor)] or LB agar [LB broth amended with 15 g L-1 Bacto agar (BD)] at 37 °C amended with 30 μg mL-1 kanamycin (LB kan) and strains containing plasmids from the

ASKA library were maintained on LB amended with 10 μg mL-1 chloramphenicol (LB cm). Plasmid DNA from ASKA library strains was extracted using a QIAprep Spin

Miniprep Kit (Qiagen). In complementation experiments with the ASKA plasmids, LB plates and top agar were supplemented with 0.05 - 0.1 mM IPTG to induce protein expression (Kitagawa et al., 2005).

Phage isolation and culture

The phages LL5 and LL12 were isolated against clinical isolates of enterotoxigenic E. coli (ETEC) obtained from John Deaton (Deerland Enzymes, Kennesaw, GA). Phages

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were isolated by the enrichment method (Gill et al., 2012) from filter-sterilized (0.22

μm) wastewater influent collected in College Station, TX in 2011. Both phages were subsequently cultured using E. coli strain DH5α as host. Phage lysates were prepared by the confluent plate lysis method (Adams, 1959) using LB (Miller) bottom plates (10 g L-

1 Bacto tryptone, 5 g L-1 Bacto yeast extract, 10 g L-1 NaCl, 15 g L-1 Bacto agar) and top agar consisting of 10 g L-1 tryptone, 10 g L-1 NaCl, 5 g L-1 Bacto agar. Phages were harvested and stored as filter-sterilized (0.22 µm) lysates in lambda diluent (25 mM Tris-

HCl pH 7.5, 100 mM NaCl, 8 mM MgSO4, 0.01% w/v gelatin) at 4 ºC (Green &

Sambrook, 2012).

Plaque assays were conducted using both spot titer and full-plate titration methods

(Adams, 1959). For spot titers, phages were diluted ten-folds and 10 μL of each dilution was spotted on solidified lawns of 4 ml top agar inoculated with 100 μL of a fresh overnight host culture prepared as described above. For full-plate titers, 100 μL of phages diluted ten-folds were mixed with 100 μL of host culture in 4 ml of molten top agar and poured over LB plates as described above. Plaques were enumerated after 16-

18 h incubation at 37 °C. The efficiency of plating (EOP) was calculated as the ratio of the number of plaques appearing on the lawn of a test strain to the number of plaques on the reference strain.

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Phage genome sequencing and annotation

Phage DNA was purified from high-titer lysates by a modified Wizard DNA purification kit (Promega) as previously described (Summer, 2009). Phage LL5 was sequenced by

454 pyrosequencing at the Emory GRA Genome Center (Emory University, GA); trimmed FLX Titanium sequence reads were assembled into a single contig at 19.9-fold coverage using Newbler 2.5.3 (454 Life Sciences) at default settings. Phage LL12 was sequenced by Illumina TruSeq as unpaired 100-base reads; reads were quality-controlled by FastQC (https://www.bioinformatics.babraham.ac.uk/projects/fastqc/) and assembled with Velvet 1.1 (Zerbino, 2010) into a single contig at 28.3-fold coverage. Assembled phage contigs were confirmed to be complete by PCR using primers facing off each end of the contig and sequencing of the resulting products. Structural annotation was conducted using Glimmer3 (Delcher et al., 1999) and MetaGeneAnnotator (Noguchi et al., 2008) with tRNAs predicted by ARAGORN (Laslett & Canback, 2004) or tRNAscan-SE (Lowe & Chan, 2016) and gene functions predicted by InterProScan

(Jones et al., 2014) or Conserved Domain Database (Marchler-Bauer et al., 2017),

TMHMM (http://www.cbs.dtu.dk/services/TMHMM), BLASTp (Camacho et al., 2009) and HHpred (Soding et al., 2005). Phage genome annotation was conducted using the

Phage Galaxy instance hosted by the Center for Phage Technology at Texas A&M

University (cpt.tamu.edu). The annotated phage genomes were deposited in NCBI

Genbank under accession no. MH491968 (LL5) and MH491969 (LL12).

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Transmission electron microscopy

Phages were stained with 2% uranyl acetate and imaged in a JEOL 1200EX transmission electron microscope (TEM) under 100 kV accelerating voltage at the Texas A&M

University Microscopy and Imaging Center, as previously described (Valentine et al.,

1968; Piya et al., 2017). The size parameters of phages were measured electronically using ImageJ (Schneider et al., 2012).

Screening and confirmation of phage-insensitive mutants

In order to optimize the input phage concentrations and incubation times, phages LL5 and LL12 stocks were diluted ten-folds in fresh LB and 160 µL of each dilution was aliquoted into 96-well sterile transparent polystyrene flat-bottom plates (Greiner Bio- one). The plates were then inoculated with the Keio parental strain BW25115 using a

96-pin replicator (Phenix) and incubated at 37 °C for 6, 8, 10 and 18 hrs. The optical density (OD) at 550 nm was measured in a Tecan M200 plate reader at each time interval and the average OD was analyzed to determine the lowest phage concentration that inhibited bacterial growth.

The Keio collection consists of 90, 96-well plates containing two independently- generated sets of 3,985 single-gene knockouts in the E. coli BW25113 background

(Baba et al., 2006). The Keio strains were replicated into 96 well sterile polypropylene

U-bottom microplates (Greiner Bio-one) containing LB kan + 8% glycerol using sterile plastic 96-pin replicators (Phenix). Plates were incubated at 37 ºC overnight and stored

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frozen at -80 ºC. These plates were used as the working stocks for the following screens.

The odd- and even-numbered plates have identical gene deletion mutants created by independent experiments (Baba et al., 2006), and only the odd-numbered 45 plates were used for the initial screen. Initial screens were conducted in 96-well sterile transparent polystyrene flat-bottom plates (Greiner Bio-one). Phages LL5 and LL12 were diluted in fresh LB to obtain working stocks of 106 PFU/mL for LL5 and 103 PFU/mL for LL12.

160 µL of the phage working stocks were aliquoted into all wells, Keio strains were inoculated into the phage lysates from the 96-well working stocks with 96-pin

° replicators, and the plates were incubated for 8 hours at 37 C. The OD550 was measured and the wells with OD550 higher than the pre-determined cutoff values (0.2 for phage

LL5 and 0.11 for phage LL12) were scored as positive for growth.

The positive mutants obtained from the first screen were verified by repeating the assay with the same strains and their corresponding mutant strains from the even-numbered

Keio collection plates, side-by-side with eight replicates per assay. Mutants that returned mean OD550 above the designated cutoff in either the even- or odd-numbered set were retained for further characterization by measurement of phage efficiency of plating (EOP) by spot assays on soft agar lawns (Adams, 1959) as described above.

EOP was calculated as the number of plaques observed on the mutant strain divided by the number of plaques observed on the parental E. coli strain BW25113. Mutants with

EOPs of less than 10-2 were confirmed by enumerating plaques on full plates. When possible, mutant alleles were moved into the parental BW25113 background by P1

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transduction using the kanamycin resistance cassette as the selectable marker (Thomason et al., 2007). All gene disruptions were confirmed by PCR using primers flanking the predicted insert followed by sequencing of the PCR product to confirm disruption of the gene. All mutants were complemented by transforming the original Keio mutant or its

P1 transductant with a plasmid expressing the corresponding gene under control of the lac promoter; all complementing plasmids were obtained from the ASKA collection

(NIG, Japan).

Results and Discussion

Isolation and characterization of phages LL5 and LL12

Phages LL5 and LL12 were isolated from municipal wastewater in College Station, TX by enrichment against enterotoxigenic E. coli (ETEC) clinical isolates. Shortly after isolation, both phages were determined to plate efficiently on E. coli K-12 strains including MG1655, DH5α and the Keio parental strain BW25113 and these phages were subsequently propagated on E. coli DH5α for the remainder of the study.

Phages LL5 and LL12 have distinct morphology as observed by transmission electron microscopy (Fig. 4.1). LL5 is a siphophage with a head diameter of 61 nm (± 2 nm) and a flexible tail 156 nm (± 9.9 nm) in length. Phage LL12 is a large myophage with head diameter of 86.4 nm (± 2.4 nm) and a tail 112.3 nm (± 4.1 nm) in length with a pronounced baseplate.

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Fig. 4.1. Transmission electron micrographs of phages LL5 and LL12. The grids for imaging were prepared as described in materials and methods. Panel A: Phage LL5 has a capsid diameter of 61 nm (±2 nm) and a tail length of 156 nm (±9.9 nm). Panel B: Phage LL12 has a capsid diameter of 86 nm (±2.4 nm) and a tail length of 112 nm (±4.1 nm). These size parameters are an average of ten measurements and the error represents standard deviation. The scale bar represents 100 nm.

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LL5 has a genome of 49,788 bp with 88 predicted protein-coding genes and no tRNA genes. The genome produced a circular assembly and was reopened be syntenic with other T1-like phages in the NCBI database such as TLS (NC_009540) and T1

(NC_005833). Thirty-three LL5 encoded proteins could be assigned putative functions

(Table 4.1, Fig. 4.2A). Genes responsible for different stages of phage infection cycle have been identified in LL5, including a DNA primase/helicase (gp58), ATP-dependent helicase (gp60) and helicase (gp75). Structural proteins including the portal protein

(gp31), major capsid protein (gp36), minor tail proteins (gp41, gp42, gp47, gp48), tail tube protein (gp43), tape measure protein (gp46), and tail fiber proteins (gp51, gp57) were identified. The small and large terminase subunits were identified as gp29 and gp30, respectively. Like its T1-like relatives, LL5 encodes a canonical lysis cassette composed of a holin (gp70), endolysin (gp71) and unimolecular spanin (gp72).

Phage LL5 is closely related to the T1-like coliphage TLS (NC_009540) (German &

Misra, 2001), with 96% sequence identity over 90% query coverage of the LL5 genome based on BLASTn analysis. As would be expected given this close similarity, the LL5 genome is syntenic with TLS, with 75 LL5 proteins having homologs in the TLS genome detectable by BLASTp with an E value of less than 10-5. Phage LL5 encodes two predicted tail fiber proteins, gp51 and gp57, in a genomic arrangement similar to that found in phage TLS and T1. LL5 gp51 is closely related to predicted tail fibers in other phages including gp51 of TLS (also called TspJ, YP_001285540, 98% identity), gp33 of phage T1 (FibA, YP_003912, 67% identity) and the central tail fiber protein J of

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Fig. 4.2. Genome maps of phages LL5 and LL12. The predicted genes of phages LL5 (panel A) and LL12 (panel B) are represented as boxes, which are numbered to match locus number. The genes present in positive and negative DNA strands of phage genomes are separated by a line; the boxes above the line denote genes in the positive strand, whereas those below the line denote genes in the negative strand. Some of the predicted gene products playing important roles in phage infection cycle are indicated. The colored boxes represent genes predicted to perform similar functions. The scale bar represents the genomic loci in kilobases.

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Fig. 4.2. Continued.

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Table 4.1. Proteins encoded by genome of phage LL5 Gene Start End Strand Gene product CPT_LL5_01 365 652 + Conserved hypothetical protein CPT_LL5_02 664 1107 + Conserved hypothetical protein CPT_LL5_03 1104 1211 + Conserved hypothetical protein CPT_LL5_04 1224 1346 - Conserved hypothetical protein CPT_LL5_05 1415 1678 + Conserved hypothetical protein CPT_LL5_06 1830 2396 + Conserved hypothetical protein CPT_LL5_07 2456 2770 + Conserved hypothetical protein CPT_LL5_08 2843 2950 + Conserved hypothetical protein CPT_LL5_09 2954 3496 + Polynucleotide kinase/phosphatase CPT_LL5_10 3493 3669 + Conserved hypothetical protein CPT_LL5_11 3764 3973 + Hypothetical protein CPT_LL5_12 4047 4301 + Conserved hypothetical protein CPT_LL5_13 4298 4480 + Conserved hypothetical protein CPT_LL5_14 4473 5048 + Conserved hypothetical protein CPT_LL5_15 5140 5340 + Conserved hypothetical protein CPT_LL5_16 5342 5506 + Conserved hypothetical protein CPT_LL5_17 5508 5633 + Conserved hypothetical protein CPT_LL5_18 5626 5871 + Conserved hypothetical protein CPT_LL5_19 5883 6353 + Putative GntR-family transcriptional regulator CPT_LL5_20 6427 6630 + Conserved hypothetical protein CPT_LL5_21 6634 6939 + Conserved hypothetical protein CPT_LL5_22 6936 7253 + Conserved hypothetical protein CPT_LL5_23 7320 7448 + Conserved hypothetical protein CPT_LL5_24 7450 7677 + Conserved hypothetical protein CPT_LL5_25 7860 8069 + Conserved hypothetical protein CPT_LL5_26 8059 8220 + Conserved hypothetical protein CPT_LL5_27 8204 8386 + Conserved hypothetical protein CPT_LL5_28 8386 8616 + Conserved hypothetical protein CPT_LL5_29 8704 9228 + Terminase small subunit CPT_LL5_30 9240 10811 + Terminase large subunit

149

Table 4.1. Continued. Gene Start End Strand Gene product CPT_LL5_31 10865 12151 + Portal protein CPT_LL5_32 12156 12827 + Conserved hypothetical protein CPT_LL5_33 12824 13933 + putative scaffold or prohead protease CPT_LL5_34 13946 14425 + Conserved hypothetical protein CPT_LL5_35 14469 14909 + Conserved hypothetical protein CPT_LL5_36 14999 15973 + Major capsid protein CPT_LL5_37 16035 16307 + Conserved hypothetical protein CPT_LL5_38 16354 16773 + Conserved hypothetical protein CPT_LL5_39 16770 17141 + Conserved hypothetical protein CPT_LL5_40 17134 17574 + Conserved hypothetical protein CPT_LL5_41 17564 17857 + Minor tail protein CPT_LL5_42 17884 18087 + Minor tail protein CPT_LL5_43 18102 18764 + tail tube protein CPT_LL5_44 18842 19156 + tapemeasure chaperone protein CPT_LL5_45 18842 19473 + tapemeasure chaperone protein frameshift product CPT_LL5_46 19511 22420 + tail tape measure protein CPT_LL5_47 22420 22767 + Minor tail protein CPT_LL5_48 22835 23593 + Minor tail protein CPT_LL5_49 23590 24312 + Tail tip assembly protein CPT_LL5_50 24305 24904 + Tail assembly protein CPT_LL5_51 24986 28762 + Tail fiber protein CPT_LL5_52 29018 29161 + Hypothetical protein CPT_LL5_53 29224 30279 + Exodeoxyribonuclease VIII CPT_LL5_54 30357 30647 + Conserved hypothetical protein CPT_LL5_55 30694 31356 + Recombinase CPT_LL5_56 31395 31820 + Single-stranded DNA binding protein CPT_LL5_57 31853 34363 - Tail fiber protein CPT_LL5_58 34495 35421 - DNA primase/helicase CPT_LL5_59 35479 36087 - Putative transcriptional regulator CPT_LL5_60 36176 38149 + ATP-dependent helicase

150

Table 4.1. Continued. Gene Start End Stran Gene product d CPT_LL5_61 38152 38559 + VRR-NUC domain protein CPT_LL5_62 38549 38683 - Conserved hypothetical protein CPT_LL5_63 38631 38909 + Conserved hypothetical protein CPT_LL5_64 38911 39651 + Dam methylase CPT_LL5_65 39653 39883 + Conserved hypothetical protein CPT_LL5_66 39922 40119 + Conserved hypothetical protein CPT_LL5_67 40201 40341 + Conserved hypothetical protein CPT_LL5_68 40344 40823 + HNH endonuclease CPT_LL5_69 40907 42022 + Conserved hypothetical protein CPT_LL5_70 42152 42358 + Putative holin CPT_LL5_71 42358 42798 + glycoside hydrolase endolysin CPT_LL5_72 42845 43249 + unimolecular spanin protein CPT_LL5_73 43266 43460 - Conserved hypothetical protein CPT_LL5_74 43384 43776 - Conserved hypothetical protein CPT_LL5_75 43779 45359 - Helicase CPT_LL5_76 45427 45675 - Conserved hypothetical protein CPT_LL5_77 45672 46061 - Conserved hypothetical protein CPT_LL5_78 46063 46257 - Hypothetical protein CPT_LL5_79 46318 47013 - Site-specific DNA methylase CPT_LL5_80 47209 47448 - Conserved hypothetical protein CPT_LL5_81 47454 47675 - Conserved hypothetical protein CPT_LL5_82 47734 47901 - Conserved hypothetical protein CPT_LL5_83 48003 48134 + Hypothetical protein CPT_LL5_84 48131 48400 - Conserved hypothetical protein CPT_LL5_85 48428 48700 + Hypothetical protein CPT_LL5_86 48684 48878 - Conserved hypothetical protein CPT_LL5_87 48875 49078 - Conserved hypothetical protein CPT_LL5_88 49150 49428 - Conserved hypothetical protein

151

phage lambda (NP_040600, 23% identity). Approximately 3 kb downstream and on the opposite strand from gp51, gp57 is similar to other T1-like tail fibers only in its N- terminal domain, with 46% identity to T1 FibB (YP_003919) from residues 1-290. The

C-terminal domain of gp57 is more closely related to tail fiber proteins found in T5-like phages such as DT57C and DT571/2 (Golomidova et al., 2015). LL5 gp57 is 57% identical with a C-proximal region spanning residues 515 to 830 of the 1,076-residue

DT57C LtfA protein (YP_009149889), which is within the host specificity region of this protein (Golomidova et al., 2016).

The LL12 genome was determined to be 136,026 bp in length and encodes 213 predicted protein-coding genes and 7 tRNAs. The genome produced a circular assembly and was reopened at a point between the genes encoding T4 RIIA and RIIB homologs, to retain its general synteny with other RIIAB-encoding myophages. Analysis of the raw

Illumina reads by PhageTerm (Garneau et al., 2017) suggests the presence of a non- permuted terminal redundancy of 459 bp spanning bases 104,966 - 105,424 in the genome as presented here. This predicted terminal repeat is located in a non-coding region of DNA between two convergent transcripts and corresponds to the location of non-permuted terminal repeats observed in phage phi92 (Schwarzer et al., 2012). Fifty

LL12-encoded proteins could be assigned putative functions, as shown in Table 4.2 and

Fig. 4.2B. Major components for head morphogenesis including capsid protein (gp59), prohead protease (gp62), portal protein (gp63), and the large terminase subunit (TerL, gp64) were identified. The components for tail morphogenesis including baseplate

152

Table 4.2. Proteins encoded by genome of phage LL12 Gene Start End Strand Gene product CPT_LL12_001 1 1983 - rIIA protector from prophage-induced early lysis CPT_LL12_002 1980 2480 - Hypothetical conserved protein CPT_LL12_003 2530 2910 - Hypothetical conserved protein CPT_LL12_004 2920 4104 - MoxR ATPase CPT_LL12_005 4104 4373 - Hypothetical conserved protein CPT_LL12_006 4418 4798 - Hypothetical conserved protein CPT_LL12_007 4801 5058 - Hypothetical conserved protein CPT_LL12_008 5058 5414 - Hypothetical conserved protein CPT_LL12_009 5428 5700 - Hypothetical conserved protein CPT_LL12_010 5703 6602 - Putative alpha 1-3 fucosyltransferase CPT_LL12_011 6745 7722 + Anti-sigma factor CPT_LL12_012 7756 9216 + Putative metallopeptidase CPT_LL12_013 9232 9495 + Hypothetical conserved protein CPT_LL12_014 9495 9758 + Hypothetical conserved protein CPT_LL12_015 9758 10021 + Hypothetical conserved protein CPT_LL12_016 10023 10256 + Hypothetical conserved protein CPT_LL12_017 10272 10721 + Hypothetical conserved protein CPT_LL12_018 10718 10963 + Hypothetical conserved protein CPT_LL12_019 10966 11532 + Hypothetical conserved protein CPT_LL12_020 11588 11863 + Hypothetical conserved protein CPT_LL12_021 11866 12339 + Hypothetical conserved protein CPT_LL12_022 12336 12506 + Hypothetical conserved protein CPT_LL12_023 12537 12917 + Hypothetical conserved protein CPT_LL12_024 13081 13488 + Hypothetical conserved protein CPT_LL12_025 13492 13692 + Hypothetical conserved protein CPT_LL12_026 13740 19376 - Hypothetical conserved protein CPT_LL12_027 19425 21170 - Tail fiber protein CPT_LL12_028 21180 21446 - Hypothetical conserved protein CPT_LL12_029 21458 22501 - Tail fiber protein CPT_LL12_030 22591 22905 - Hypothetical conserved protein

153

Table 4.2. Continued Gene Start End Strand Gene product CPT_LL12_031 22917 23501 - Tail fiber assembly protein CPT_LL12_032 23516 24556 - Tail fiber protein CPT_LL12_033 24569 26641 - Tail fiber protein CPT_LL12_034 26641 27327 - Hypothetical conserved protein CPT_LL12_035 27339 28829 - Baseplate protein CPT_LL12_036 28935 32534 - Tail fiber protein CPT_LL12_037 32534 32959 - Hypothetical conserved protein CPT_LL12_038 32959 33510 - Hypothetical conserved protein CPT_LL12_039 33512 34102 - Hypothetical conserved protein CPT_LL12_040 34200 34532 - Hypothetical conserved protein CPT_LL12_041 34532 38971 - Tail fiber protein CPT_LL12_042 39007 41604 - Tail fiber protein CPT_LL12_043 41606 42274 - Hypothetical conserved protein CPT_LL12_044 42284 43000 - Hypothetical conserved protein CPT_LL12_045 43013 43690 - Hypothetical conserved protein CPT_LL12_046 43690 44694 - Hypothetical conserved protein CPT_LL12_047 44694 45074 - Hypothetical conserved protein CPT_LL12_048 45086 45970 - Hypothetical conserved protein CPT_LL12_049 46075 48414 - Hypothetical conserved protein CPT_LL12_050 48469 48717 - Hypothetical conserved protein CPT_LL12_051 48729 49202 - Hypothetical conserved protein CPT_LL12_052 49366 49839 - Hypothetical conserved protein CPT_LL12_053 49850 51226 - Tail sheath protein CPT_LL12_054 51299 51850 - Hypothetical conserved protein CPT_LL12_055 51850 52275 - Hypothetical conserved protein CPT_LL12_056 52291 52749 - Hypothetical conserved protein CPT_LL12_057 52803 53423 - Hypothetical conserved protein CPT_LL12_058 53483 54307 - Hypothetical conserved protein CPT_LL12_059 54392 55399 - Major capsid protein CPT_LL12_060 55448 55837 - Head stabilization/decoration protein

154

Table 4.2. Continued Gene Start End Strand Gene product CPT_LL12_061 55858 56835 - Hypothetical conserved protein CPT_LL12_062 56835 57254 - Prohead protease CPT_LL12_063 57322 58878 - Portal protein CPT_LL12_064 58981 60847 - Terminase large subunit CPT_LL12_065 60847 61134 - O-spanin CPT_LL12_066 61131 61529 - I-spanin CPT_LL12_067 61635 61904 - Hypothetical conserved protein CPT_LL12_075 62861 63199 - Conserved hypothetical protein CPT_LL12_076 63451 63561 - Conserved hypothetical protein CPT_LL12_077 63546 63788 - Conserved hypothetical protein CPT_LL12_078 63865 64053 - Conserved hypothetical protein CPT_LL12_079 64064 64642 - Conserved hypothetical protein CPT_LL12_080 65303 65653 - Putative transcriptional regulator CPT_LL12_081 65701 66255 - Phosphoesterase CPT_LL12_082 66263 66673 - ATP-binding protein CPT_LL12_083 66792 67709 + RNA ligase and tail attachment protein CPT_LL12_084 67706 68056 + Hypothetical conserved protein CPT_LL12_085 68065 68826 + Putative Sir2-like protein CPT_LL12_086 68840 69553 + Putative Sir2-like protein CPT_LL12_087 69616 69924 + Conserved hypothetical protein CPT_LL12_088 69911 70180 + Conserved hypothetical protein CPT_LL12_089 70180 71463 + DNA ligase CPT_LL12_090 71587 72057 + Endolysin

155

Table 4.2. Continued Gene Start End Strand Gene product CPT_LL12_091 72121 72327 + Conserved hypothetical protein CPT_LL12_092 72337 72801 + HNH endonuclease CPT_LL12_093 72794 73084 + Conserved hypothetical protein CPT_LL12_094 73144 73452 + Conserved hypothetical protein CPT_LL12_095 73445 74584 + Exodeoxyribonuclease CPT_LL12_096 74584 74916 + Conserved hypothetical protein CPT_LL12_097 74913 75542 + Hypothetical conserved protein CPT_LL12_098 75494 76099 + EndoVII packaging and recombination endonuclease CPT_LL12_099 76038 76259 + Conserved hypothetical protein CPT_LL12_100 76272 77255 + Putative DNA polymerase/exonuclease CPT_LL12_101 77301 78119 + Putative DNA N6-adenine methyltransferase CPT_LL12_102 78082 78540 + Hypothetical conserved protein CPT_LL12_103 78568 78759 + Hypothetical conserved protein CPT_LL12_104 78759 79373 + Hypothetical conserved protein CPT_LL12_105 79367 79960 + Hypothetical conserved protein CPT_LL12_106 79962 80162 + Hypothetical conserved protein CPT_LL12_107 80162 81145 + Thymidylate synthase CPT_LL12_108 81244 81705 + Hypothetical conserved protein CPT_LL12_109 81716 81946 + Hypothetical conserved protein CPT_LL12_110 81943 84258 + Ribonucleoside triphosphate reductase alpha chain CPT_LL12_111 84298 85386 + Ribonucleoside diphosphate reductase beta chain CPT_LL12_112 85390 85668 + Glutaredoxin 1 CPT_LL12_113 85665 87788 + Anaerobic ribonucleoside triphosphate reductase CPT_LL12_114 87852 87968 + Hypothetical conserved protein CPT_LL12_115 87984 88088 + Hypothetical protein CPT_LL12_116 88120 88356 + Hypothetical conserved protein CPT_LL12_117 88353 88826 + Anaerobic ribonucleoside triphosphate reductase activating protein CPT_LL12_118 88883 88996 + Hypothetical conserved protein CPT_LL12_119 88993 89343 + Hypothetical conserved protein CPT_LL12_120 89382 90170 + PhoH-like protein

156

Table 4.2. Continued Gene Start End Strand Gene product CPT_LL12_121 90269 90580 + Hypothetical conserved protein CPT_LL12_122 90640 91689 + Clp ATP-dependent protease subunit CPT_LL12_123 91741 92274 + DNA methyltransferase CPT_LL12_124 92271 92555 + Hypothetical conserved protein CPT_LL12_125 92590 92829 + Hypothetical conserved protein CPT_LL12_126 92826 93224 + Hypothetical conserved protein CPT_LL12_127 93264 93497 + Hypothetical conserved protein CPT_LL12_128 93503 94027 + Hypothetical conserved protein CPT_LL12_129 94058 94294 + Hypothetical conserved protein CPT_LL12_130 94278 94430 + Hypothetical conserved protein CPT_LL12_131 94514 94699 + Hypothetical conserved protein CPT_LL12_132 94702 95157 + Hypothetical conserved protein CPT_LL12_133 95218 95565 + Hypothetical conserved protein CPT_LL12_134 95562 95849 + Hypothetical conserved protein CPT_LL12_135 95846 96040 + Hypothetical conserved protein CPT_LL12_136 96060 96245 + Hypothetical conserved protein CPT_LL12_137 96341 96637 + Hypothetical conserved protein CPT_LL12_138 96640 96921 + Hypothetical conserved protein CPT_LL12_139 96921 97145 + Hypothetical conserved protein CPT_LL12_140 97135 97389 + Hypothetical conserved protein CPT_LL12_141 97402 97800 + Hypothetical conserved protein CPT_LL12_142 97818 98108 + Hypothetical conserved protein CPT_LL12_143 98191 98508 + Sigma 54 modulation factor CPT_LL12_144 98646 98915 + Hypothetical conserved protein CPT_LL12_145 99007 99270 + Hypothetical conserved protein CPT_LL12_146 99275 99607 + Hypothetical conserved protein CPT_LL12_147 99607 100002 + Hypothetical conserved protein CPT_LL12_148 100082 100339 + Hypothetical conserved protein CPT_LL12_149 100340 100690 + Hypothetical conserved protein CPT_LL12_150 100690 101019 + Hypothetical conserved protein

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Table 4.2. Continued Gene Start End Strand Gene product CPT_LL12_151 101019 101306 + Hypothetical conserved protein CPT_LL12_152 101396 101659 + Hypothetical conserved protein CPT_LL12_153 101688 101828 + Hypothetical conserved protein CPT_LL12_154 101815 102066 + Hypothetical conserved protein CPT_LL12_155 102095 102400 + Hypothetical conserved protein CPT_LL12_156 102453 102779 + Hypothetical conserved protein CPT_LL12_157 102788 103087 + Hypothetical conserved protein CPT_LL12_158 103355 103660 + Hypothetical conserved protein CPT_LL12_159 103782 103997 + Hypothetical conserved protein CPT_LL12_160 106517 106936 - Hypothetical conserved protein CPT_LL12_161 107082 107345 - Hypothetical conserved protein CPT_LL12_162 107406 107678 - Hypothetical conserved protein CPT_LL12_163 107897 108208 - Hypothetical conserved protein CPT_LL12_164 108205 108396 - Hypothetical conserved protein CPT_LL12_165 108413 108703 - Hypothetical conserved protein CPT_LL12_166 108795 109010 - Hypothetical conserved protein CPT_LL12_167 109020 109124 - Hypothetical conserved protein CPT_LL12_168 109202 109447 - Hypothetical conserved protein CPT_LL12_169 109540 109719 - Hypothetical conserved protein CPT_LL12_170 109795 110028 - Hypothetical conserved protein CPT_LL12_171 110083 110367 - Hypothetical conserved protein CPT_LL12_172 110570 110920 - Hypothetical conserved protein CPT_LL12_173 110938 111237 - Hypothetical conserved protein CPT_LL12_174 111331 111531 - Hypothetical conserved protein CPT_LL12_175 111679 111798 - Hypothetical conserved protein CPT_LL12_176 111819 112058 - Hypothetical conserved protein CPT_LL12_177 112018 112389 - Hypothetical conserved protein CPT_LL12_178 112454 112834 - Hypothetical conserved protein CPT_LL12_179 112923 113546 - Hypothetical conserved protein CPT_LL12_180 113622 113882 - Hypothetical conserved protein

158

Table 4.2. Continued Gene Start End Strand Gene product CPT_LL12_181 114102 114299 - Hypothetical conserved protein CPT_LL12_182 114289 114642 - Hypothetical conserved protein CPT_LL12_183 114717 114962 - Hypothetical conserved protein CPT_LL12_184 115052 115393 - Hypothetical conserved protein CPT_LL12_185 115473 115592 - Hypothetical conserved protein CPT_LL12_186 115673 116068 - Hypothetical conserved protein CPT_LL12_187 116151 116549 - Hypothetical conserved protein CPT_LL12_188 116614 116811 - Hypothetical conserved protein CPT_LL12_189 117664 117810 + Hypothetical conserved protein CPT_LL12_190 117913 118242 - Hypothetical conserved protein CPT_LL12_191 118239 118433 - Hypothetical conserved protein CPT_LL12_192 118426 118671 - Hypothetical conserved protein CPT_LL12_193 118681 119013 - Hypothetical conserved protein CPT_LL12_194 119023 119214 - Hypothetical conserved protein CPT_LL12_195 119278 119472 - Hypothetical conserved protein CPT_LL12_196 119469 119669 - Hypothetical conserved protein CPT_LL12_197 119723 120181 - Hypothetical conserved protein CPT_LL12_198 120171 120596 - Hypothetical conserved protein CPT_LL12_199 120665 120862 - Hypothetical conserved protein CPT_LL12_200 120958 121374 - Hypothetical conserved protein CPT_LL12_201 121437 122663 - Hypothetical conserved protein CPT_LL12_202 122663 123028 - Hypothetical conserved protein CPT_LL12_203 123112 123357 - Hypothetical conserved protein CPT_LL12_204 123375 123677 - Hypothetical conserved protein CPT_LL12_205 123667 124056 - Hypothetical conserved protein CPT_LL12_206 124056 124328 - Hypothetical conserved protein CPT_LL12_207 124392 124646 - Hypothetical conserved protein CPT_LL12_208 124723 125094 - Hypothetical conserved protein CPT_LL12_209 125105 125299 - Hypothetical conserved protein CPT_LL12_210 125296 125523 - Hypothetical conserved protein

159

Table 4.2. Continued Gene Start End Strand Gene product CPT_LL12_211 125520 125825 - Hypothetical conserved protein CPT_LL12_212 125854 128907 - DNA polymerase A CPT_LL12_213 128966 131044 - DNA replicative helicase/primase CPT_LL12_214 131034 131798 - DNA cytosine methyltransferase CPT_LL12_215 131850 132332 - Hypothetical conserved protein CPT_LL12_216 132345 132740 - Hypothetical conserved protein CPT_LL12_217 132740 133042 - Hypothetical conserved protein CPT_LL12_218 133103 133444 - Hypothetical conserved protein CPT_LL12_219 133441 134811 - Helicase CPT_LL12_220 134811 136016 - rIIB-like protein

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(gp35), tail sheath protein (gp53), and multiple predicted tail fiber proteins (gp27, gp29, gp32, gp33, gp36, gp41, gp42) were identified. DNA replication proteins such as DNA polymerase (gp212), DNA replicative helicase/primase (gp213), and a helicase (gp219) were also identified. The genes encoding the large terminase subunit (gp64) and DNA polymerase (gp212) are disrupted by predicted intron sequences. These introns appear to be relatively short (~275-325 bp) and do not contain any significant protein-coding

ORFs. The boundaries of these introns were determined based on protein sequence similarity to homologous proteins found in other phages that were not disrupted by introns (AKU44155 in the case of gp64, and AKU44295 for gp212). Like other large myophages, genes responsible for phage lysis are distributed across the LL12 genome rather than co-localized to a contiguous cassette; the phage endolysin (gp90), i-spanin

(gp66) and o-spanin (gp65) were identifiable but the phage holin could not be positively identified. Based on analysis of predicted protein sequences by BLASTp with a E value cutoff of 10-5, LL12 is most closely related to other V5-like myophages, including rV5

(NC_011041) which shares 206 proteins with LL12, and ΦAPCEc02 (KR698074) which shares 204 proteins. LL12 is also more distantly related to the E. coli phage phi92

(NC_023693), with 48 common proteins detectable by BLASTp.

Like the related phages rV5, phi92 and ΦAPCEc02, phage LL12 encodes an extensive set of predicted tail fibers: gp27, gp29, gp32, gp33, gp36, gp41 and gp42 (Fig. 4.2B,

Table 4.2). All seven of these LL12 tail fibers are similar to the tail fibers in found in rV5 and ΦAPCEc02, with protein identities ranging from 43%-100% (Table 4.3). Six of

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Table 4.3. Analysis of LL12 tail fibers LL12 BlastP hits tail % of % E- alignment fibers identical identity Phage Protein Gene value quality locus Phage matches (Dice) tag Accession # Accession # product rV5 NC_011041.1 YP_002003530.1 gp028 99.0 0 0.99 CPT_ APCEc02 KR698074.1 AKO61946.1 98.0 0 0.98 weak, LL12_ phi92 NC_023693.1 YP_009012483.1 gp151 47.0 7E-23 0.08 027 partial weak, phi92 NC_023693.1 YP_009012482.1 gp150 43.9 2E-22 0.07 partial

APCEc02 KR698074.1 AKO61944.1 99.1 0 0.99 CPT_ rV5 NC_011041.1 YP_002003532.1 gp30 98.9 0 0.99 LL12_ rV5 NC_011041.1 YP_002003535.1 gp33 43.0 2E-90 0.44 029 APCEc02 KR698074.1 AKO61941.1 43.0 2E-90 0.44 phi92 NC_023693.1 YP_009012479.1 gp147 23.5 6E-18 0.25 full length

APCEc02 KR698074.1 AKO61941.1 99.4 0 0.99 rV5 NC_011041.1 YP_002003535.1 gp33 99.1 0 0.99 CPT_ APCEc02 KR698074.1 AKO61944.1 43.3 6E-91 0.44 LL12_ rV5 NC_011041.1 YP_002003532.1 gp30 42.7 1E-88 0.44 032 N- and C- phi92 NC_023693.1 YP_009012479.1 gp147 27.0 9E-29 0.30 terminus only

rV5 NC_011041.1 YP_002003536.1 gp34 99.9 0 1.00 CPT_ APCEc02 KR698074.1 AKO61940.1 99.7 0 1.00 LL12_ good, 033 phi92 NC_023693.1 YP_009012474.1 gp142 42.2 0 0.41 almost full length

CPT_ APCEc02 KR698074.1 AKO61937.1 99.0 0 0.99 LL12_ 036 rV5 NC_011041.1 YP_002003539.1 gp37 83.6 0 0.86

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Table 4.3. Continued

LL12 BlastP hits tail % of % gene E- alignmen fibers Phage Protein identical identity Phage product value t quality locus Accession # Accession # matches (Dice) tag APCEc02 KR698074.1 AKO61932.1 95.9 0 0.96 rV5 NC_011041.1 YP_002003543.1 gp41 91.5 0 0.70 CPT_ rV5 NC_011041.1 YP_002003545.1 gp43 38.4 3E-15 0.04 LL12_ 041 APCEc02 KR698074.1 AKO61930.1 38.4 3E-15 0.04 very phi92 NC_023693.1 YP_009012483.1 gp151 37.7 3E-07 0.02 weak, patchy

rV5 NC_011041.1 YP_002003544.1 gp42 99.4 0 0.99 CPT_ APCEc02 KR698074.1 AKO61931.1 99.2 0 0.99 LL12_ 042 good, phi92 NC_023693.1 YP_009012473.1 gp141 34.95 2E-173 0.36 almost full length

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these seven proteins are also detectable in the more distantly-related phage phi92, with three of these, gp29, gp33 and gp42 producing alignments to nearly the full-length phi92 proteins 147, 142 and 141, respectively (Table 4.3). CryoEM reconstructions of phi92 have indicated that this phage possesses multiple sets of tail fibers that are mounted to the baseplate in downward, sideward, and upward orientations (Schwarzer et al., 2012).

These multiple tail fibers may contribute to a broadened host range in this phage and its relatives (Schwarzer et al., 2012). The electron density of the downward-facing tail fiber was assigned to gp143, which is not conserved in LL12 (Schwarzer et al., 2012).

LL12 gp27 shows weak similarity to the N-terminus of phi92 gp150, which is predicted to form downward-facing tail spikes in cryoEM reconstructions (Schwarzer et al., 2012).

LL12 gp41 also possesses similarity to rV5 gp41 (Table 4.3), however LL12 gp41 is missing the C-terminal chaperone of endosialidase domain (pfam13884) of rV5 gp41 spanning residues 1151-1200.

Host range determination for phages LL5 and LL12

Infection by STEC strains can result in watery or bloody diarrhea, hemolytic uremic syndrome, microangiopathic hemolytic anemia and thrombocytopenia (Gyles, 2007). E. coli strains belonging to several pathotypes tend to be clonal and are grouped as serotypes based on the O-antigens (lipopolysaccharide) and H-antigens (flagella) (Kaper et al., 2004). As phage LL12 bears similarity to phages rV5 and ΦAPCEc02, both of which infect STEC serotype O157:H7 (Kropinski et al., 2013; Dalmasso et al., 2016),

164

we sought to determine if phages LL5 and LL12 are also able to infect STEC representatives.

Phages LL5 and LL12 were spotted on soft agar overlays of STEC strains and EOP compared to the Keio parental strain BW25113. Phage LL5 was unable to form plaques on any of the tested STEC strains, and phage LL12 exhibited EOPs of close to 1 on

STEC strains with serotypes O157:H7, O145:NM, O121:H19, O111 and O121:H19, demonstrating a relatively broad host range among STEC serotypes for this phage (Table

4.4).

Development and optimization of screening assay

Multiplicity of Infection (MOI) is the ratio of the number of the phages to host cells in a culture. The purpose of the phage Keio screen was to identify host genes required for the phage to successfully infect, replicate within, and lyse their host cells. To determine this, it was imperative to optimize MOI for each phage as excessively high MOI’s could result in bacterial growth inhibition if the phage were able to infect the cells but still not produce progeny, while too low MOI’s could result in false positive results

(Georgopoulos, 2006). Initially, the lowest input phage concentration required to control growth of parental BW25113 strain after 8 hr incubation at 37 °C was determined. A log higher phage concentration was applied in this screen so as to minimize false positives.

The number of bacterial cells inoculated by the 96-pin replicator were determined by

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Table 4.4. Host range of phages LL5 and LL12. Phage LL5 and LL12 were tested for their ability to infect Shiga toxin-producing Escherichia coli (STEC) by spotting serially diluted phages on the soft agar lawns of respective STEC isolates. The efficiency of plating (EOP) is relative to the number of plaques formed on the Keio collection parental E. coli strain BW25113. Cells marked with "-" indicate an EOP of less than 10-7 (insensitive to phage). The data is the average of two biological replicates. STEC serotype Isolate ID Phage LL5 Phage LL12 LPS Core EOP EOP typesb Not STEC BW25113 1.0 1.0 K-12 O157:H7a USDA-FSIS 380-94 - 0.8 R3 O104:H21 ATCC BAA-178 - - O145:NMa 83-75 - 0.7 R1, K-12 O26:H11a H30 - - R3 O111:H-a JBI-95 - - R3 O121:H19 ATCC BAA-2219 - 0.7 O146 ATCC BAA-2217 - 1.0 O103:H11 ATCC BAA-2215 - - O145:Nonmotile ATCC BAA-2192 - - R1, K-12 O26:H11 ATCC BAA-2196 - - R3 O45:H2 ATCC BAA-2193 - - O103:H2a CDC 90-3128 - - R3 O121:H19a CDC 97-3068 - 0.6 O45:H2a CDC 96-3285 - - aSources of these isolates are described in (Kirsch et al., 2014). bLPS core types information obtained from (Amor et al., 2000).

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viable counts. Based upon the cells inoculated and PFU of phages used, the initial MOI of LL5 and LL12 in this screen was calculated to be 1.0 and 0.001, respectively.

Determination of genes required for phage propagation

The Keio collection consists of a total of 3,985 individual gene knockout mutants in the

E. coli K-12 strain BW25113. Each gene knockout is represented twice in the collection

(the results of two independent experiments) (Baba et al., 2006), thus the total collection contains 7,970 mutants, with each independent gene knockout mutants represented with even and odd numbers. Phages LL5 and LL12 were screened against the entire odd- numbered series of 3,985 Keio single-gene knockouts as described above. E. coli mutants that were unable to support phage growth, as indicated by their growth to an

OD550 of at least 0.2 or 0.11 at 8 h in the presence of phage LL5 or LL12, respectively, were considered positive hits in this initial screen. Using this selection criteria, 37 knockout mutants (21 mutants for each phage) were selected for further investigation

(Tables 4.5, 4.6, 4.7). For each of these initial hits, the screening experiment was repeated using the same odd-numbered mutant and its even-numbered counterpart from the collection. From this second experiment, 11/21 mutants identified against LL5 and

9/21 mutants identified against LL12 were found to produce the same phenotype in at least one of the paired knockouts, and these were retained for further study (Tables 4.5,

4.6, 4.7).

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Table 4.5. Summary of hits in Keio screening. Initial screening of the Keio collection against both phages LL5 and LL12 gave 21 hits. These initial hits were verified using both of the independently generated Keio mutants using the same screening procedure. For phages LL5 and LL12, 10 and 9 strains, respectively, were determined to be true positives, which were tested for plating efficiency of the respective phages. Phage LL5 had a reduced efficiency of plating (EOP) of at least ~10-2 , as determined by spot titer assay, in 9 strains. Only 6 of these strains showed similar plating defect in full plate assay. Only 4 of these strains could be P1 transduced and phage LL5 had a similar plating defect, as compared to the original Keio mutants, in 3 of the P1 transductants. Phage LL12 had a reduced EOP of at least ~10-2 in 4 strains, as determined by spot titer. All 4 of these strains showed similar plating defect in full plate assay. Only 1 of these strains could be P1 transduced and phage LL12 had similar plating defect in the P1 transductant, as compared to the original Keio strain. Whenever applicable, gene complementation to restore the plating defect was conducted on the P1 transduced strains. When the kanamycin resistance marker could not be transduced, gene complementation was assayed on the original Keio mutants.

Strains with plating defect P1 Initial Verified P1 transduced strain Phages transduced hits hits with plating defect strains Spot titer Full plate titer

LL5 21 10 8 4 4 3 LL12 21 9 3 3 1 1

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Table 4.6. Results of initial (untargeted) screening and targeted re-screening of phage LL5 against the Keio E. coli knockout collection. In the initial screen, all mutants yielding a positive result were screened a second time against both independently-generated gene knockouts present in the Keio collection, denoted as the representatives from the even- and odd-numbered plate sets. Mutants with a positive result from either set were then tested for their efficiency of plating (EOP) by both spot titer and full-plate titration methods. Mutants exhibiting a significant EOP defect (less than ~0.05) were used for further study. The presence of the appropriate gene deletion was confirmed by PCR and sequencing. One knockout from each even/odd pair was selected for P1 transduction of the kan-marked deletion into the parental E. coli strain BW25113 background, re-tested for EOP defects and complemented in trans. Selected gene knockouts from the collection that were not identified in the initial screen were targeted for re-screening (bottom panel). Mutants were cultured from the Keio collection, the presence of the appropriate gene deletion confirmed by PCR, and the EOP determined in the knockout and its complemented counterpart. Blank cells denote that data was not collected, usually because the desired EOP defects were not observed.

Initial screen of Keio collection Results of second EOP (spot titer) EOP (plate titer) Keio EOP Results mutant Deletion EOP (plate (plate Gene screen Transducible of first used for confirmed titer) in P1 titer) in deletion by P1vir screen further by PCR transductant compleme- Odd Even Odd Even Odd Even work nted strain

nuoM + + + 0.06 0.04 0.3 0.3 yehQ + - - ycdB + + - 0.4 0.2 ydcR + + - 1 1 ygcN + - -

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Table 4.6. Continued. Results of Keio second EOP (spot titer) EOP (plate titer) EOP (plate Results mutant Deletion EOP (plate Gene screen Transducible titer) in of first used for confirmed titer) in P1 deletion by P1vir compleme- screen further by PCR transductant Odd Even Odd Even Odd Even nted strain work rof + - - crcB + - - ydjO + - - holD + - - aroC + - - ydgL + - - idi + - - rfaP + + + < 8 x 10-8 1.0 < 7.5 x 10-8 1.1 Odd Confirmed Yes < 7.5 x 10-8 2.2 rfaG + + - 1.1 0.3 Confirmed rfaF + + + 0.3 0.03 1.3 0.3 rfaQ + + + 0.01 0.4 0.04 0.04 Odd Confirmed Yes 0.3 rfaH + - + 0.003 0.01 0.6 0.2 rfaY + - - secB + + + 0.01 0.03 0.05 0.02 Odd Confirmed Yes 0.06 0.2 ppiB + + - 0.003 0.3 0.04 0.4 Odd Confirmed Yes 0.09 1.5 ylaC + + - 0.07 0.03 0.4 0.8

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Table 4.6 Continued. Targeted re-screening Gene Deletion Keio mutant EOP (spot EOP (plate EOP (plate titer) in deletion confirmed by used for further titer) titer) complemented strain PCR work lpcA Confirmed Even < 8 x 10-8 < 7.5 x 10-8 0.8 rfaE Confirmed Even < 8 x 10-8 < 7.5 x 10-8 1.0 rfaC Loci intact

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Table 4.7. Results of initial (untargeted) screening and targeted re-screening of phage LL12 against the Keio E. coli knockout collection. In the initial screen, all mutants yielding a positive result were screened a second time against both independently-generated gene knockouts present in the Keio collection, denoted as the representatives from the even- and odd-numbered plate sets. Mutants with a positive result from either set were then tested for their efficiency of plating (EOP) by both spot titer and full-plate titration methods. Mutants exhibiting a significant EOP defect (less than ~0.05) were used for further study. The presence of the appropriate gene deletion was confirmed by PCR and sequencing. One knockout from each even/odd pair was selected for P1 transduction of the kan-marked deletion into the parental E. coli strain BW25113 background, re-tested for EOP defects and complemented in trans. Selected gene knockouts from the collection that were not identified in the initial screen were targeted for re-screening (bottom panel). Mutants were cultured from the Keio collection, the presence of the appropriate gene deletion confirmed by PCR, and the EOP determined in the knockout and its complemented counterpart. Blank cells denote that data was not collected, usually because the desired EOP defects were not observed.

Initial screen of Keio collection

Results of Keio second EOP (spot titer) EOP (plate titer) EOP (plate Gene Results mutant Deletion EOP (plate screen Transducible titer) in deletio of first used for confirmed titer) in P1 by P1vir complemen- n screen Odd Even Odd Even Odd Even further by PCR transductant ted strain work pflC + - - cusB + - - ompR + + - 0.5 0.5

envZ + - - ompC + + + 0.3 0.3 yncJ + - -

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Table 4.7 Continued. Results of Keio second EOP (spot titer) EOP (plate titer) EOP (plate Results mutant Deletion EOP (plate Gene screen Transducible titer) in of first used for confirmed titer) in P1 deletion by P1vir complemen- screen further by PCR transductant ted strain Odd Even Odd Even Odd Even work

ycbL + - - yqjB + nt nt 1.9 1.1

yaaW + - - yggT + + + 0.1 0.1

rfaP + + + 0.06 1.1 0.04 1.2 Odd Confirmed Yes 0.02 1.0 rfaD + - - rfaG + + - 7.1 x 1.0 5.1 x 1.3 Odd Confirmed No 1.1 10-6 10-6 rfaF + - - rfaH + + - 0.4 0.8

rfaY + + + 0.3 0.2 rffE + + - 0.2 0.1

lpcA + - + 0.8 < 4 x 1.5 < 4.4 x Even Confirmed No 1.1 10-9 10-9 trmC + - - ygbF + - - etp + - -

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Table 4.7 Continued Targeted re-screening Deletion Keio mutant Gene EOP (spot EOP (plate EOP (plate titer) in confirmed by used for deletion titer) titer) complemented strain PCR further work rfaE Confirmed Even < 4 x 10-9 < 4.4 x 10-9 1.1 rfaI Confirmed Odd 0.6 rfaB Confirmed Even 0.4

rfaC Loci intact rfaF Loci intact

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The efficiency of plating (EOP) of phage LL5 on the retained mutant strains was determined by spot titer. The observed plating efficiency of phage LL5 was reduced by at least ~20-fold in eight mutants. This plating defect was confirmed by titration of LL5 in full plate assays, in which only four mutants showed an EOP reduction of ~20-fold or greater. In order to confirm the plating phenotype in a clean genetic background, the kanamycin resistance cassettes from these Keio mutants were transduced by P1 into the parental E. coli strain BW25113. Markers could be transduced from all four Keio mutants into the parental strain, and three showed a similar plating defect as the corresponding Keio mutant, indicating the phenotype was linked to the disrupted locus

(Table 4.5). One mutant, rfaQ, showed a ~25-fold reduction in EOP in the Keio mutant but its P1 transductant exhibited only a very mild EOP defect of 0.3 (Table 4.6), despite having the rfaQ deletion confirmed in the transductant by PCR and sequencing. This suggests an abnormality or additional defect in the original rfaQ Keio mutant; this mutant was not examined further.

The same approach was applied to confirm the phenotypes of the Keio mutants identified from the screens against phage LL12. Spot titer assays showed that the EOP of phage LL12 was reduced in only three of the nine initially identified Keio mutants. This plating defect could be replicated via full plate plaque assay in all three mutants. Only one of these mutants could be P1 transduced into the parental strain BW25113 and same plating phenotype was observed in the P1 transductant as in the Keio mutant (Table 4.5).

The other two Keio mutants were resistant to P1 infection and could not be transduced.

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Based on the genes identified in these initial screens, additional mutants from the odd- and even-numbered Keio sets were subjected to targeted re-screening by directly determining the phage EOP by the spot method. For both phages, genes involved in LPS biosynthesis (rfaP for LL5, and rfaP, rfaG and lpcA for LL12, Tables 4.6, 4.7, 4.8) were identified and confirmed, but these genes represented only parts of the known biosynthetic pathway. Additional Keio mutants in lpcA, rfaE, rfaC, rfaF, rfaI and rfaB were confirmed by PCR and sequencing of the mutant locus; lpcA, rfaE, rfaI and rfaB were found to contain the appropriate deletions. Strong EOP defects (< 10-8) were identified in the lpcA and rfaE mutants against LL5, and in rfaE in LL12 (an EOP defect in lpcA against LL12 was already identified in the initial screen) (Table 4.8). These mutants could not be transduced due to P1 resistance.

Five genes were determined to be required for efficient propagation of phage LL5, as evident by the reduced plating efficiency (Table 4.8). The Keio mutant strains deleted for genes rfaP, lpcA and rfaE showed severe plating defects with EOP of phage LL5 less than 8 x 10-8. This plating defect was also observed on the P1 transduced rfaP mutant.

The plating efficiency of phage LL5 could be restored when the respective genes were provided in trans (Table 4.8). All plating phenotypes were complemented in the P1 transduced strains when applicable, and in the original Keio strains when the mutations could not be transduced by P1 due to host resistance. Two additional Keio mutants, secB and ppiB, exhibited milder defects in supporting phage LL5 growth, with EOP

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Table 4.8. E. coli genes required for phages LL5 and LL12 propagation. The genes required for phage infection cycle can be determined by testing the efficiency of plating. Five genes were found to be required for phage LL5 infection cycle, whereas 4 genes were required for phage LL12 infection cycle. The kanamycin resistance cassette in the Keio strain was P1 transduced into parental BW25113, whenever possible. The P1 transductants are denoted by "#" and the original Keio mutants are denoted by "*". The plating phenotype was complemented in P1 transductants, when applicable. The data represents average and standard deviation of three biological repeats. Phages Gene EOP Complemented EOP lpcA* < 8 x 10-8 0.8 ± 0.2 rfaE* < 8 x 10-8 1.0 ± 0.5 LL5 rfaP# < 8 x 10-8 2.2 ± 2.0 secB# 0.06 ± 0.02 0.2 ± 0.04 ppiB# 0.09 ± 0.05 1.5 ± 0.3 lpcA* < 4.4 x 10-9 1.1 ± 0.4 rfaE* < 4.4 x 10-9 1.1 ± 0.5 LL12 rfaP# 0.02 ± 0.01 1.0 ± 0.4 rfaG* 5.1 x 10-6 ± 1.0 x 10-6 1.1 ± 0.2

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reductions of ~10- to 100-fold relative to the parental E. coli strain BW25113; these mutants could be transduced by P1 into the parental background and could also be complemented in trans (Table 4.8).

The three genes lpcA, rfaE and rfaP are parts of the LPS biosynthesis pathway. The severe plating defects associated with multiple genes in this pathway strongly indicate that phage LL5 uses LPS as its receptor, and LPS defects result in major blocks to phage infection. Of these three genes, only rfaP appeared as a “hit” in the initial screen of the

3,985 Keio mutants against phage LL5 (Table 4.6). RfaP adds phosphate or 2- aminoethyl diphosphate (PPEtN) to heptose (Hep) I of the inner core LPS, and the LPS of rfaP mutants does not have any phosphoryl substituents on Hep I or Hep II, and also lacks Hep III (Fig. 4.3) (Yethon et al., 1998). Hep I is transferred to LPS from the nucleotide precursor molecule ADP-L-glycero-D-manno-heptose, which is synthesized in a separate pathway involving lpcA, rfaE and rfaD (Fig. 4.3) (Gronow & Brade, 2001).

In the absence of either LpcA or RfaE, heptoseless LPS core is formed (Valvano et al.,

2000), whereas rfaD mutants can still incorporate the stereoisomer D-glycero-D-manno- heptose into the LPS (Coleman & Leive, 1979). Keio lpcA and rfaE mutants with heptoseless LPS core cannot be infected by bacteriophage P1, rendering P1 transduction of the genetic marker ineffective (Valvano et al., 2000).

As the rfaP mutant is the last step in the LPS pathway with a strong observed plating defect for LL5, this raises a question about the role of Hep II phosphorylation and the

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Fig. 4.3. Genes and biosynthetic pathway of the E. coli lipopolysaccharide (LPS) required for replication of phages LL5 and LL12. Proteins in the pathway are denoted in bold and label each step in biosynthesis. Black triangles indicate proteins that are required by LL5 for growth, and black circles indicate proteins required by LL12. Panel A: The nucleotide sugar precursor ADP L-glycero-β-D-manno-heptose is used as a substrate for the transfer of heptose (green in panel B) to the E. coli core LPS. ADP L-glycero-β-D-manno-heptose is synthesized from sedoheptulose-7-P via a pathway comprised of LpcA, RfaE and RfaD. Both LpcA and RfaE are required for growth of phages LL5 and LL12. Panel B: LPS is composed of four distinct domains: Lipid A, inner core, outer core and O-antigen. The enzymes responsible for the addition of sugar residues and phosphoryl constituents relevant to this study are denoted. RfaC, RfaF, RfaQ, RfaG, RfaI and RfaB add hexo or hepto sugar residues to LPS, and RfaP and RfaY add phosphoryl substituents to the heptose residues I and II, respectively. RfaP is required for infection by phage LL5 and the open circle by RfaP indicates a milder plating defect (~50-fold reduction in plating efficiency) for phage LL12 in the absence of this protein. RfaG is required for LL12 infection. Panel A is adapted from (Gronow & Brade, 2001) and panel B from (Clifton et al., 2013).

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addition of Hep III in the LL5 host recognition mechanism. RfaY adds the phosphoryl group to Hep II and RfaQ links Hep III to Hep II (Yethon et al., 1998). The Keio rfaY mutant was ruled out during the initial screen and the plating defect of phage LL5 on the rfaQ transductant was mild (Table 4.6). From this information, it can be inferred that the plating defect of phage LL5 on the rfaP mutant is solely due to the absence of phosphoryl substituents on Hep I. Moreover, no other molecule is linked to the phosphoryl group of Hep I, thus it can be concluded that inner core of LPS is used by phage LL5 as its receptor and the phosphoryl group of Hep I is required by phage LL5 to infect its E. coli host. The LPS receptor requirement of phage LL5 appears similar to those of phage TLS (German & Misra, 2001). Phage TLS utilizes outer membrane protein TolC and the sugar residues Hep I (and its phosphoryl group), and Hep II of the

LPS inner core to recognize and infect its E. coli host (German & Misra, 2001). The putative tail fibers of LL5 bear similarity to those of TLS, so it was intriguing why tolC did not come up as in screen. The requirement of tolC for LL5 infection still needs to be tested.

Apart from the three genes that were involved in LPS biosynthesis, the plating efficiency of phage LL5 was also reduced in Keio strains with deletion in two other genes, secB and ppiB. The EOP of phage LL5 was 0.06 and 0.09 in secB and ppiB strains, respectively. The plating defect in both strains could be restored to normal levels when the gene was supplied in trans (Table 4.8). Both SecB and PpiB are chaperones that

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contribute to protein translocation and proline peptide bond isomerization, respectively

(Bechtluft et al., 2010; Unal & Steinert, 2014).

Proteins, once synthesized in cytoplasm, are sorted into compartments of the cell by different protein transport systems. SecB, a tetrameric cytoplasmic chaperone, is a component of the general secretory (Sec) system that transports proteins synthesized in the cytoplasm, post-translationally, to the extra-cytoplasmic compartments. Post- translational transport is primarily preferred for periplasmic and outer membrane proteins (Denks et al., 2014; Findik & Randall, 2017). SecB binds to polypeptides and keeps them in an unfolded state until cytoplasmic ATPase SecA directs the bound polypeptide to the SecYEG transmembrane channel (Findik & Randall, 2017). The translocation of polypeptides across the SecYEG channel is powered by the cytoplasmic

ATPase SecA (Denks et al., 2014). Eighteen E. coli proteins have been reported to be dependent on SecB-mediated translocation (Bechtluft et al., 2010) (Findik & Randall,

2017). We do not know if any of these SecB-dependent proteins play a role in the infection cycle of phage LL5, or if the potential accumulation of cytoplasmic protein aggregates in secB mutants hampers phage replication. In the absence of SecB, other cytoplasmic chaperones have been reported to be upregulated to stabilize secretory proteins during their delayed translocation and/or to rescue protein aggregates (Baars et al., 2006). This compensatory mechanism by other chaperones may be the reason why the EOP defect of phage LL5 in the secB mutant is relatively mild (EOP = 0.06).

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Another chaperone affecting the plating efficiency of phage LL5 is PpiB, which belongs to peptidyl-prolyl cis/trans isomerase (PPIase) superfamily of proteins, catalyzing protein folding at the peptide bonds preceding proline residues (Unal & Steinert, 2014).

Although PPIases play a role in several biological processes, there is no evidence of any biological process depending solely on any PPIases (Unal & Steinert, 2014). The genome of E. coli K-12 encodes eight PPIases, belonging to three families: FKPBs, cyclophilins and parvulins (Unal & Steinert, 2014). The cyclophilins family consist of

PpiA and PpiB, which are periplasmic and cytoplasmic proteins, respectively (Unal &

Steinert, 2014). To our knowledge, there is only one reported instance of the requirement of a PPIase for a phage infection cycle: SlyD, belonging to FKBP family of PPIases, has been shown to be required for plaque formation by the ssDNA phage ΦX174 (Roof et al., 1994). SlyD is required to stabilize ΦX174 lysis protein E, so that it can accumulate to optimum levels to lyse the host cell (Bernhardt et al., 2002). Since the infection cycle of phage LL5 has not been characterized, it is difficult to explain which aspect of phage replication is affected by the absence of PpiB.

Four genes were determined to be required for efficient propagation of phage LL12, as evident by the plating defect (Table 4.8). All Keio strains in which phage LL12 showed plating defects were deleted for genes in the LPS biosynthesis pathway. Phage LL12 showed severe plating defects (EOP < 10-8) in lpcA and rfaE deletions, and an EOP of

~10-6 in the rfaG deletion. The plating defect of phage LL12 was milder (EOP ~0.02) in the rfaP deletion (Table 4.8). The plating defects of phage LL12 could be restored when

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the respective genes were supplied in trans. All complementation assays were conducted in the P1 transduced strains when applicable, and in the original Keio strains when the mutations could not be transduced by P1 (Table 3).

The functions of genes lpcA, rfaE and rfaP in LPS biosynthesis have been explained in context of phage LL5 above. RfaG links glucose (Glc) I to Hep II of the LPS inner core

(Fig. 4.3) (Parker et al., 1992; Yethon et al., 2000), and marks the start of the outer core domain of the E. coli LPS. Sugar residues Glc II and galactose (Gal) are linked to Glc I by RfaI and RfaB respectively (Schnaitman & Klena, 1993). The plating efficiency of phage LL12 in the respective rfaI and rfaB mutants were close to wild type (~0.5) suggesting that Glc II and the Gal sidechain do not play significant roles in phage LL12 infection (Table 4.7). The strongly reduced EOP of phage LL12 on rfaG deletions suggests a crucial role of the outer core Glc I in the host recognition mechanism of phage LL12. This Hep II - Glc I linkage is conserved in K-12, and R1 - R4 LPS core types in E. coli (Amor et al., 2000). As shown in Table 4.4, phage LL12 is able to infect

E. coli strains with K-12, R1 and R3 LPS core types, which is consistent with the finding that the Gal sidechain residue linked to Glc I in the K-12 core and the residues downstream of Glc I are not involved in phage receptor binding. To our knowledge,

LL12 is the first candidate from the group of V5-like phages for which the host receptor has been characterized. Based upon the sequence similarities of their tail fibers, other closely related V5-like phages such as rV5, ΦAPCEc02, and the O157:H7 typing phages

5 and 14 are likely to use the same or similar receptors as phage LL12.

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From genetic analysis, we have established that phage LL12 recognizes the E. coli LPS core as its host receptor in a K-12 background. However, LL12 is also able to infect multiple different serotypes of E. coli with varying O-antigen (Table 4.4). In the case of phage P1, the extensive O-antigen expressed by hosts such as E. coli O157:H7 and

Salmonella Typhimurium is able to obscure the LPS core and loss of the O-antigen results in bacterial sensitivity to this phage (Ornellas & Stocker, 1974; Ho & Waldor,

2007). This observation suggests that phage LL12 has developed a mechanism to deal with the presence of O-antigen that may obscure its receptor in the LPS core. Several phages are known to have evolved mechanisms to reach the cell surface to recognize these polysaccharide coats to facilitate infection. The tail spike protein (TSP) of

Salmonella phage P22 recognizes O-antigen as its receptor and also has endorhamnosidase activity and cleaves its glycosidic linkages resulting in the shortening of the O-antigen (Andres et al., 2010a; Andres et al., 2010b). Coliphage G7C also expresses tail spikes with enzymatic activity against O-antigen that is involved in phage adsorption (Prokhorov et al., 2017).

Conclusions

The Keio collection is a library of single-gene deletion of non-essential genes in E. coli

K-12 strain BW25113. The library consists of 7970 mutants, with each mutant generated independently and distributed in odd- and even-numbered plates (Baba et al., 2006).

Phages LL5 and LL12 were initially isolated against pathogenic E. coli hosts as

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candidates for therapeutic use, and phage LL12 was shown to infect representatives of several prominent STEC serovars. Both phages LL5 and LL12 were screened against the

Keio library to investigate the host factors required for successful phage propagation.

Initial screens suggested that twenty-one E. coli genes were necessary for phage LL5 and LL12 propagation (Table 4.5), but on further analysis a total of 5 and 3 E. coli genes were found to affect the propagation of phages LL5 and LL12, respectively (Tables 4.5,

4.8). Based on these figures, the screening process resulted in a false positive rate of

86%; the false negative rate of this screen cannot be calculated but target re-testing of genes of interest identified two additional genes not identified in the initial screens.

These observations highlight the generally noisy nature of high-throughput screens and the requirement for additional confirmatory experiments following screening.

Through successive verification screen, spot and full plate plaque assays, it was established that three E. coli genes were needed for each of phage LL5 and LL12 propagation (Table 4.8), which gives a false positive rate of ~86% for both phages. Of the genes required for propagation of either phage LL5 or LL12, ~60% of the genes constitute the LPS biosynthesis pathway. It was noteworthy that all the genes pertaining to the LPS pathway did not come in the initial screen “hits”, so targeted screen by spot- titer method was done for phage LL5 against Keio mutants lpcA and rfaE, and for phage

LL12 against Keio mutant rfaE (Table 4.8); the gene disruption in Keio mutants lpcA and rfaE was verified beforehand. Both phages LL5 and LL12 exhibited plating defects on the respective Keio mutants.

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The genes presented in this study may not be exhaustive list of genes necessary for phage LL5 or LL12 propagation because of limitations of these high throughput screens.

If any biological pathway is comprised of multiple genes, it is likely that some genes will appear as “hits” in the initial screen and the role of other genes of that particular pathway can be tested by using verified gene mutants. The Keio collection, nevertheless, is a great library for initial screen, but data should be interpreted only after exhaustive follow-up of the mutants.

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CHAPTER V

CONCLUSIONS AND FUTURE DIRECTIONS

Co-evolution of phages and their bacterial hosts have resulted in development of several defense and counter-defense strategies (Chapter I). This dissertation focuses on the study of antirestriction system of coliphage P1, employed to overcome bacterial type I restriction and modification (R-M) systems (Chapters II and III). Two components of P1 antirestriction system, DarA and DarB, were described previously (Iida et al., 1987).

From genetic analysis, it was demonstrated that DarA protects P1 DNA from type I

EcoA, whereas DarB protects P1 DNA from type I EcoB and EcoK systems (Iida et al.,

1987). Chapter II of this dissertation provides evidence suggesting P1 antirestriction is comprised of other components besides DarA and DarB. It has been shown that DarB and Ulx are required for protection against EcoB and EcoK systems, whereas Hdf,

DarA, and DdrA are required for protection against EcoA, EcoB and EcoK systems.

Interestingly, disruption of ddrB provided increased protection against EcoB and EcoK systems. Biochemical evidence supported by genetic analysis, suggests that components of P1 antirestriction system are incorporated into P1 virions in a definite order. Hdf and

DarA are incorporated first, followed by DdrA. DarB, Ulx and DdrB are incorporated next. Specific order of incorporation of DarB, Ulx and DdrB is not known, but it seems that Ulx can be incorporated only in the presence of DarB (Chapter II). The role of Hdf and DarA in capsid morphogenesis has also been demonstrated in Chapter II. When either hdf or darA is disrupted, ~80% of the virion progenies have aberrant small heads.

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The roles played by Hdf and DarA in head-size determination and incorporation of antirestriction proteins suggest that P1 antirestriction system is linked to capsid morphogenesis.

Chapter III further explores P1 antirestriction system following upon information provided in Chapter II. Since the P1 antirestriction components were encoded in two operons separated by genes of unrelated functions, it was hypothesized that other regions of P1 could encode for other unknown components of the P1 antirestriction system. To determine if there are any other components of P1 antirestriction system, isogenic knockouts of P1 genes of unknown functions, were constructed. Out of 29 genes deleted,

24 were found not to be required for P1 propagation at tested laboratory conditions.

Isogenic P1 mutants disrupted for these 24 genes were tested for restriction phenotype by plating assays. None of these 24 genes were found to be required for antirestriction function. However, five genes pmgA, pmgB, pmgC, pmgG and pmgR were found to be required for P1 propagation. The essentiality of these genes still need to be confirmed by in trans complementation. After the phenotype of these genes have been confirmed, lysates of these mutants can be observed under transmission electron microscopy to see if these mutants have any morphogenesis defect. Specific roles of these genes in virion morphogenesis, if any, can then be further studied.

To understand the mechanism of antirestriction, incorporation of DarB into P1 capsids was further studied (Chapter III). Evidence presented in Chapter III suggests that P1

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antirestriction components are packaged before DNA. It has been demonstrated in

Chapter II that proteins of P1 antirestriction system contribute to a significant mass in P1 capsids. Since P1ΔdarA cannot package any antirestriction components, it could have greater internal volume compared to that of P1. As DNA is packaged into P1 procapsids by headful mechanism (Bachi & Arber, 1977; Lobocka et al., 2004), it can be hypothesized that P1ΔdarA can package more DNA because of the availability of more internal capsid volume. This can be tested by pulsed-field gel electrophoresis of phage

DNA (Lingohr et al., 2009).

SDS-PAGE analysis of procapsid/tail fractions of P1ΔpacA and P1ΔdarA_pacA lysates indicate presence of novel proteins in lanes of both P1ΔpacA and P1ΔdarA_pacA, which are missing in the lane of CsCl purified P1 (Chapter III). Proteins present in procapsids, but not in mature virions, are indicative of scaffolding core (Dokland, 1999). These novel proteins can be identified by mass spectrometry and if determined to be virion- associated, their role in capsid morphogenesis can be further explored.

Moreover, evidence suggests that incorporation of DarB into procapsid is guided a signal provided by N-terminal residues of DarB (Chapter III). By testing incorporation of recombinant foreign proteins fused at their N-terminus with N-terminal DarB residues into P1 virions, it can be determined if these residues are sufficient for incorporation.

Moreover, it can also be determined if these residues are sufficient for ejection of proteins into host following infection. In this study, the first 9 and 30 N-terminal

189

residues of DarB have been fused to TetR-mCherry fusion protein. By testing the incorporation of these fusion proteins into P1ΔdarB, the necessity and sufficiency of these selected N-terminal DarB residues for capsid targeting can be determined.

Even though genetic evidence has established the role of DarB in protecting P1 DNA from host restriction, biochemical activity of protection still needs to be determined in vitro. As protein purification protocol for DarB has been optimized, further biochemical assay can be conducted with purified DarB to elucidate biochemical mechanisms for its antirestriction activity. Since DarB can be incorporated into P1 capsids only in the presence of other P1 proteins Hdf, DarA, DdrA, it is possible that DarB is interacting directly with some of these proteins. Also, DarB might be interacting with host components such as type I R-M subunits and host chaperones for its activity. Since the affinity tag for DarB purification has been optimized, same tag can be exploited to determine interacting partners of DarB by conducting pull-down assays.

High throughput genetic screens were conducted for novel T1-like coliphage LL5 and rV5-like coliphage LL12 against the E. coli Keio collection to understand components of phage-host interactions. Receptors for both phages LL5 and LL12 were characterized and two chaperones, PpiB and SecB were demonstrated to be required for efficient propagation of phage LL5. Bioinformatic analysis suggests that both phages encode multiples sets of tail fibers. The receptor binding proteins of both phage LL5 and LL12

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have not been characterized. Further experiments can be designed to characterize the receptor binding proteins of phages LL5 and LL12.

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